Hideyuki Tanabe* † , Stefan Mu¨ ller*, Michaela Neusser*, Johann von Hase ‡ , Enzo Calcagno ‡ , Marion Cremer*, Irina Solovei*, Christoph Cremer ‡ , and Thomas Cremer* ^@

*Department of Biology II—Human Genetics, University of Munich, Richard Wagner Strasse 10, 80333 Mu¨nchen, Germany;
† Cell Bank Laboratory, Division of Genetics and Mutagenesis, National Institute of Health Sciences, 1-18-1, Kamiyoga, Setagaya-ku, Tokyo 158-8501, Japan; and
‡ Kirchhoff Institute of Physics, University of Heidelberg, Albert Ueberle Strasse 3-5, 69120 Heidelberg, Germany

^@ To whom reprint requests should be sent. E-mail: thomas.cremer@lrz.uni-muenchen.de

We demonstrate that the nuclear topological arrangement of chromosome territories (CTs) has been conserved during primate evolution over a period of about 30 million years. Recent evidence shows that the positioning of chromatin in human lymphocyte nuclei is correlated with gene density. For example, human chromosome
19 territories, which contain mainly gene-dense and early replicating chromatin, are located toward the nuclear center, whereas chromosome 18 territories, which consist mainly of gene-poor and later replicating chromatin, is located close to the nuclear border. In this study, we subjected seven different primate species to comparative analysis of the radial distribution pattern of human chromosome 18- and 19-homologous chromatin by three-dimensional fluorescence in situ hybridization. Our data demonstrate that gene-density-correlated radial chromatin arrangements were conserved during higher-primate genome evolution, irrespective of the major karyotypic rearrangements that occurred in different phylogenetic lineages. The evolutionarily conserved positioning of homologous chromosomes or chromosome segments in related species supports evidence for a functionally relevant higher-order chromatin arrangement that is correlated with gene-density.

Introduction:

The chromatin of individual chromosomes is organized in chromosome territories (CTs) that are essential components of the higher-order chromatin architecture of the vertebrate cell nucleus (reviewed in refs. 1–5). Recently, the extent to which evolutionarily conserved, cell type, cell cycle, and species-specific motifs of chromatin arrangements may exist, has become the focus of intense studies (6–9). In mammals, two principal
components of mitotic chromosomes can be distinguished: G-light bands (also called R-bands) replicate early during S-phase and contain most of the housekeeping but relatively few tissue-specific genes. G-dark bands replicate later, are gene poor, and contain tissue-specific genes (10). Recently it has been demonstrated that these chromosome bands are maintained in interphase nuclei as focal chromatin aggregations (11) built up by a
number of chromatin domains in the order of ~1 Mb. These domains apparently persist through all interphase stages, show distinct nuclear localization patterns, and may provide an important component of the higher-order nuclear architecture (refs. 11–14, for review see ref. 2). The nuclear location of human (Homo sapiens, HSA) chromosomes 18 and 19 CTs (further referred to as HSA18 and HSA19) has become of special interest
in this respect. These chromosomes are of similar DNA content (86 Mb and 72 Mb, respectively; ref. 15) but differ strongly in their gene content and replication timing: most of HSA19 chromatin belongs to G-light bands, is gene-dense [20.5 genes per megabase pair (Mbp)], whereas most of HSA18 chromatin represents G-dark bands and consists mainly of gene-poor chromatin (4.3 genes per Mbp; ref. 10 and
http://www.ensembl.org/Homo_sapiens/). In human lymphocyte nuclei, which exhibit an almost spherical shape, HSA19 CTs are consistently localized toward the nuclear center without any detectable attachment to the nuclear envelope, whereas the HSA18 CTs are positioned close to the nuclear border (7, 8). In this study we
demonstrate the evolutionary conservation of radial nuclear arrangements for chromosomes or chromosome segments homologous to HSA18 and 19 in seven higher-primate species. The last common ancestor of these species dates back approximately 30–40 million years ago. This evolutionary conservation argues
for a still unknown functional significance of distinct radial higher-order chromatin arrangements.
...
Discussion:

Our results demonstrate that the distinctly different radial distribution patterns that have been found for CTs 18 and 19 in human lymphocyte and lymphoblastoid cell nuclei have been conserved for HSA18- and HSA19-homologous chromatin during higher-primate evolution. In all species analyzed, HSA18-homologous chromatin was found at the nuclear periphery and HSA19-homologous chromatin was found toward the nuclear
interior. This radial distribution pattern was thus maintained over a period of at least 30 million of years, irrespective of the extensive chromosomal rearrangements that occurred during the evolution of higher primates. Our results fit the hypothesis that radial chromatin arrangements reflect differences in gene density
(6). Additional evidence supporting this hypothesis is provided by our observation of a specific orientation of the relatively gene-poor HSA1q32->qter and the gene-dense HSA19- homologous chromatin segment, which form chromosome 14 in the squirrel monkey. This hypothesis also holds true for somatic translocation events. For example, a somatic t(18;19) translocated chromosome also maintained the original nuclear orientation
of the translocation partners with a peripheral location of HSA18 and an internal location of the HSA19 region (8).

It is a well established fact that the positioning of genes close to heterochromatin blocks can strongly affect their transcription (25), and it has also been argued that heterochromatin blocks may play a role with regard to the evolving nuclear architecture (26). Chromosome translocations that join heterochromatic segments
with gene-dense chromatin segments may therefore lead to radial chromatin shifts depending on the size and composition of the respective segments. For example, gene-poor chromatin and heterochromatin has often been noted at the nuclear periphery (5).

Accordingly, the joining of a heterochromatin block with a gene-dense chromosome segment, previously located in a gene-dense interior nuclear compartment, may result in a positional shift of the latter toward the peripheral nuclear compartment, possibly affecting its transcriptional activity. Our present study provides two possible examples for such a shift. First, the squirrel monkey chromosome 14p (homologous to HSA1q32->qter) is heteromorphic in the cell line used in this study (Figs. 1 and 2c). One 14 p-arm shows a large additional heterochromatic band, which is neither present in the other homologous squirrel monkey chromosome 14
nor in the corresponding counterparts of marmoset chromosome 18 and tamarin chromosome 20 (Fig. 1). This additional band may possibly explain why squirrel monkey 14p chromatin was distributed on average more toward the nuclear periphery (maximum DNA content at 79%) than marmoset 18 and tamarin 20 CTs showing
their maximum DNA content at 65% and 62%, respectively (Table 2, Fig. 6). Second, we observed a more exterior position of CTs of HSA19 homologues in chimpanzee and gorilla (at relative radius of 58% and 60%, respectively) as compared with human and orangutan (at 48% and 38%, respectively; Table 2, Fig. 5). Chimpanzee and gorilla homologues contain large regions of terminal heterochromatin (Figs. 1, arrows, and 2a), whereas these blocks are absent in the respective human and orangutan chromosomes (Fig. 1). It is intriguing to look for other examples of chromosome evolution as well as chromosomally rearranged tumor cells, where the chromatin context adjacent to a given gene-poor or gene-rich chromosome segment changes, and to test the consequences for radial positioning and gene function.

Finally, the finding of specific radial CT arrangements is not limited to primate cell nuclei, but was also reported for chicken cell nuclei (9). In Gallus gallus domesticus, microchromosomes are early replicating and considerably more gene-dense than the gene-poor and later replicating macrochromosomes. We noted the location of microchromosome territories preferentially in the nuclear interior surrounded by the more peripherally located macrochromosome territories. This gene-density-correlated radial higher-order chromatin arrangement in chicken cell nuclei shows that the evolutionary conservation of nonrandom radial arrangements is compatible with drastic changes in karyotype evolution that have occurred before the separation of the
evolutionary branches that led to present days mammals and birds. In this context, it is interesting to note that syntenic regions of HSA19 have been assigned to chicken microchromosomes, whereas syntenic regions of HSA18 have been assigned to the chicken macrochromosomes 2 and Z (27, 28).

The evidence for an evolutionary conservation of gene-density-correlated radial chromatin arrangements argues for a functional significance. Possible underlying molecular mechanisms responsible for the establishment and maintenance of these higher-order chromatin arrangements remain to be elucidated. In this context, it is interesting to test whether the density of expressed genes rather than of all genes plays a major role. The observation of different positions of the active and inactive X chromosome in female cell nuclei argues for such a possibility. In addition, the different CG content of gene-dense and gene-poor chromosome segments should be considered (29).

Support:

This study was supported by a stipend of the Japanese government, Science and Technology Agency (to H.T.), Deutsche Forschungsgemeinschaft Grant Cr 59/20-1 (to T.C.), European Community Grant FIGH-CT 1999-00011 (to C.C.), and by a grant in part, Health Sciences Research Grants, Ministry of Health, Labor and Welfare, Japan.

References:

1. Chevret, E., Volpi, E. V. & Sheer, D. (2000) Cytogenet. Cell Genet. 90, 13–21.

2. Cremer, T. & Cremer, C. (2001) Nat. Rev. Genet. 2, 292–301.

3. Cremer, T., Kreth, G., Koester, H., Fink, R. H. A., Heintzmann, R., Cremer, M., Solovei, I. V., Zink, D. & Cremer, C. (2000) Crit. Rev. Eukaryotic Gene Expression 12, 179–212.

4. Lamond, A. I. & Earnshaw, W. C. (1998) Science 280, 547–553.

5. Leitch, A. R. (2000) Microbiol. Mol. Biol. Rev. 64, 138–152.

6. Boyle, S., Gilchrist, S., Bridger, J. M., Mahy, N. L., Ellis, J. A. & Bickmore, W. A. (2001) Hum. Mol. Genet. 10, 211–219.

7. Cremer, M., v. Hase, J., Volm, T., Brero, A., Kreth, G., Walter, J., Fischer, C., Solovei, I., Cremer, C. & Cremer, T. (2001) Chromosome Res. 9, 541–567.

8. Croft, J. A., Bridger, J. M., Boyle, S., Perry, P., Teague, P. & Bickmore, W. A. (1999) J. Cell Biol. 145, 1119–1131.

9. Habermann, F., Cremer, M., Walter, J., Hase, J., Bauer, K., Wienberg, J., Cremer, C., Cremer, T. & Solovei, I. (2001) Chromosome Res. 9, 569–584.

10. Craig, J. M. & Bickmore, W. A. (1994) Nat. Genet. 7, 376–382.

11. Sadoni, N., Langer, S., Fauth, C., Bernardi, G., Cremer, T., Turner, B. M. & Zink, D. (1999) J. Cell Biol. 146, 1211–1226.

12. Jackson, D. A. (1995) BioEssays 17, 587–591.

13. Nakayasu, H. & Berezney, R. (1989) J. Cell Biol. 108, 1–11.

14. O’Keefe, R. T., Henderson, S. C. & Spector, D. L. (1992) J. Cell Biol. 116, 1095–1110.

15. Lander, E. S., Linton, L. M., Birren, B., Nusbaum, C., Zody, M. C., Baldwin, J., Devon, K., Dewar, K., Doyle, M., FitzHugh, W., et al. (2001) Nature (London) 409, 860–921.

16. Mu¨ller, S., Neusser, M., O’Brien, P. C. M. & Wienberg, J. (2001) Chromosome Res. 9, 689–693.

17. Mu¨ller, S., O’Brien, P. C., Ferguson-Smith, M. A. & Wienberg, J. (1997) Hum. Genet. 101, 149–153.

18. Solovei, I., Walter, J., Cremer, M., Habermann, F., Schermelleh, L. & Cremer, T. (2002) in FISH: A Practical Approach, eds. Squire, J., Beatty, B. & Mai, S. (Oxford Univ. Press, Oxford), in press.

19. Telenius, H., Carter, N. P., Bebb, C. E., Nordenskjold, M., Ponder, P. A. I. & Tunnacliffe, A. (1992) Genomics 13, 718–725.

20. Stanyon, R., Consigliere, S., Mu¨ller, S., Morescalchi, A., Neusser, M. & Wienberg, J. (2000) Am. J. Primatol. 50, 95–107.

21. Stanyon, R., Consigliere, S., Bigoni, F., Ferguson-Smith, M., O’Brien, P. C. & Wienberg, J. (2001) Chromosome Res. 9, 97–106.

22. Wienberg, J. & Stanyon, R. (1998) ILAR J. 39, 77–91.

23. Yunis, J. J. & Prakash, O. (1982) Science 215, 1525–1530.

24. Jauch, A., Wienberg, J., Stanyon, R., Arnold, N., Tofanelli, S., Ishida, T. & Cremer, T. (1992) Proc. Natl. Acad. Sci. USA 89, 8611–8615.

25. Brown, K. E., Baxter, J., Graf, D., Merkenschlager, M. & Fisher, A. G. (1999) Mol. Cell. 3, 207–217.

26. Manuelidis, L. (1990) Science 250, 1533–1540.

27. Burt, D. W., Bruley, C., Dunn, I. C., Jones, C. T., Ramage, A., Law, A. S., Morrice, D. R., Paton, I. R., Smith, J., Windsor, D., et al. (1999) Nature (London) 402, 411–413.

28. Schmid, M., Nanda, I., Guttenbach, M., Steinlein, C., Hoehn, M., Schartl, M., Haaf, T., Weigend, S., Fries, R., Buerstedde, J. M., et al. (2000) Cytogenet. Cell Genet. 90, 169–218.

29. Bernardi, G. (2001) Gene 276, 3–13.

1. Frenster JH, "Uni-Polar Clustering of Lymphocyte DNA Templates Toward Neoplastic Target Cells Within Hodgkin's Disease Lymph Nodes".

2. Frenster JH, Papalian MM, Masek MA, and Frenster JA, "Electron Microscopic Analysis of Lymph Node Cellular Activity in Hodgkin's Disease".

3. Frenster JH, "Electron Microscopic Localization of Acridine Orange Binding to DNA within Human Leukemic Bone Marrow Cells".

4. Frenster JH, "Ultrastructural Probes of Active DNA Sites, and the RNA Activators of DNA".

5. Frenster JH, "Activation of DNA Transcription within Repressed Chromatin".

Top of Page - Euchromatin Network - Current Research - Forums - Other Sites - Future Events -