four control regions were selected: −16 kb, upstream of tal1; + 5 and + 22 kb, within the tal1 locus; + 70 kb, downstream of tal1. these sequences were chosen as control regions as they show limited homology between a number of species ( 26 ) and were considered highly unlikely to represent regulatory elements. all four regions showed no enrichments between dnasei - treated and - untreated samples in either the hes cells or k562 cells. the most striking differences between hes cells and k562 cells were found at the tal1 regulatory elements. with k562 cells, there is prominent enrichment at the tal1 promoter 1a, and the −4, + 19 and + 50 enhancers. these cells express high levels of tal1 transcripts, and were originally derived from a patient with blastic transformation of chronic myeloid leukaemia. k562 cells are relatively undifferentiated, although they have a partial erythroid phenotype. this potentially explains the marked enrichment seen at the + 19 stem cell enhancer and the + 50 erythroid enhancer. this is similar to data obtained from primary mouse erythroid cells ( day 14. 5 foetal liver ), which show the highest enrichments at the mouse + 40 enhancer ( homologous to the human + 50 ) ( data not shown ). in contrast, the hes cells show markedly reduced enrichment at the tal1 regulatory elements when compared with k562 cells. although there is enrichment at the + 50 enhancer, there is minimal enrichment at the −4, promoter 1a and + 19 elements. reduced / absent accessibility of regulatory elements to dnasei digestion is consistent with the lack of expression of tal1 in hes cells ( figure 4a ). as es cells differentiate to form embryoid bodies, expression of tal1 is rapidly switched on, appearing from day 3 of mouse es cell differentiation ( 32 ). the relative accessibility of the + 50 enhancer may reflect a poised chromatin state in hes cells, which permits a rapid response to the changing transcription factor environment that accompanies differentiation. the development of techniques that permit the rapid comparative mapping of dhs between different cell types will greatly facilitate the study of transcriptional regulation in both normal and diseased cells. recently published high - throughput techniques that map dhs sites using high - throughput sequencing ( 8, 9 ) and genomic array tiling paths ( 7, 11, 13 ) have clear advantages of scale over more targeted approaches. major disadvantages of these approaches include cost and lack of focus. this makes them less suitable for many laboratories that want to assess the chromatin accessibility of a number of defined or presumed regulatory elements in a range of cell types. a real - time - pcr - based approach to dhs mapping has, therefore, a number of potential advantages for researchers interested in specific regulatory questions at defined loci. real - time pcr is relatively inexpensive compared with the large - scale techniques, and permits a rapid, focused dhs analysis of defined regions of dna from multiple cell types. it also provides flexibility, as any genomic region can be analysed from the dna library derived from the dnasei - treated samples by designing further real - time pcr primer sets. we have previously shown that our basic technique for amplifying a library of dhs generates a template representative of known dhs with excellent sensitivity and specificity ( 11 ). although the experiments presented in this paper were each performed using 5 million cells / digestion condition, we have obtained reproducible data using 5 - fold fewer cells as starting material. we feel our technique can deliver acceptable specificity and sensitivity for dhs mapping with small numbers of cells, and will therefore be of use to those researchers working with limited numbers of primary cells. an alternative approach using real - time pcr to define dhs has been previously published ( 10 ). the two approaches differ in that dorscher et al. quantify the dhs through the loss of pcr signal obtained from dhs when dna is digested, whereas our approach uses real - time pcr to quantify a gain of signal observed from dhs. dorscher et al. report excellent sensitivity using their approach to map dhs. however, the technique depends on large numbers of comparative quantitative real - time pcr reactions across a region in both digested and undigested material, in order to quantify the loss of enrichment. one advantage of our technique published here is that data can be obtained using far fewer quantitative pcr reactions. the technique is highly reproducible, with relatively little variation in quantifiable enrichments observed between different biological replicates. moreover, we demonstrate tissue specificity, with variable enrichment at known regulatory elements between different cell types. the technique published here permits the rapid comparative analysis of dhs between different cell types from relatively small numbers of cells. it will have potential use for researchers across a broad spectrum of biology for the study of transcriptional regulation in both healthy and diseased tissues. cell culturecell lines were maintained in culture as previously described ( 11 ). care was taken to ensure maximum viability of cells when taken for experiments. the primary thymocytes used for figure 2 were prepared by gentle physical disassociation of a whole thymus gland into pbs supplemented with 2 % fcs. cells were filtered to ensure that a single - cell suspension was taken forward for nuclei isolation. human embryonic stem cells ( hes ) were grown in chemically defined media in the presence of activin and fgf2, as detailed previously ( 33 ). in these conditions, hes cells remain homogenously undifferentiated. nuclei preparationup to 3 × 107 cells were washed in ice - cold pbs and resuspended in 2 ml of ice - cold cell lysis buffer [ 300 mm sucrose, 10 mm tris ph 7. 4, 15 mm nacl, 5 mm mgcl, 0. 1 mm egta, 60 mm kcl, 0. 2 % np - 40, 0. 5 mm dtt, 0. 5 μm spermidine, 1× protease inhibitor ( complete, roche ) ]. after 5 min, the lysed cells were spun at 500 g for 5 min at 4°c with brakes off. after careful removal of supernatant, the nuclei were gently resuspended in 200 μl of ice - cold reaction buffer ( 20 μl 10× dnasei buffer, 4 μl glycerol, 176 μl water ) using pipette tips with cut off ends. the nuclei were spun again at 500 g for 5 min at 4°c and, following supernatant removal, were resuspended in 30 μl of reaction buffer per dnasei condition. for example, if 2 × 107 cells were taken to look at four different conditions ( e. g. 0, 20, 60, 120 units dnasei ), the nuclei were resuspended in 120 μl of reaction buffer. separate 30 μl aliquots were then taken and gently mixed with 70 μl of dnasei mix ( see table 1 ) on ice. this made a final digestion volume of 100 μl for each sample, which was left to incubate for 1 h on ice in the cold room. after 1 h, 700 μl of nuclei lysis buffer ( 100 mm tris hcl ph 8, 5 mm edta ph 8, 200 mm nacl, 0. 2 % sds ) was added to each sample with 50 μg proteinase k. following gentle mixing with inversion, the lysed samples were incubated at 55°c for 1 h. rnasea ( 10 μg ( ambion ) ) was then added to each sample and further incubated at 37°c for 30 min. dna was then extracted using standard phenol – chloroform techniques. care was taken to use cut - off tips and very gentle pipetting to reduce non - specific dna sheering. following precipitation, dna was resuspended in 200 μl of 0. 1 te and quantified using spectophotometry. samples ( 1 μg ) were analysed using gel - electrophoresis, as shown in figure 1a. dna library preparationfollowing quantification, 7. 5 μg of dna was taken for each sample condition. this was blunt - ended using t4 polymerase ( 10 μl 10× buffer, 0. 5 μl 25 mm dntp, 3 μl bsa, 1 μl t4 polymerase ( 3 u / μl ne biolabs ), 85. 5 μl of water / dna in 0. 1 te ). the samples were mixed gently on ice, then put at 12°c for 16 min.