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Mapping Protein/DNA Interactions by Cross-Linking [Internet]. Paris: Institut national de la santé et de la recherche médicale; 2001.

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Part I. Chromatin Protein Complexes: Dynamic Assemblies in the Nucleus

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Abstract

Our research focuses on the influence of specific repressor and activator complexes on nuclear architecture in the context of Drosophila development using a combination of fluorescence in situ hybridization, specific GFP--protein constructs, and immunofluorescence in high-resolution imaging microscopy. I will discuss the Polycomb-group repression complex, which maintains silencing of the homeobox genes in specific tissues throughout development, and the MSL dosage compensation complex, which up-regulates the level of transcription of genes on the X chromosome in males.

Nuclear Architecture and Functional Consequences

Positioning of chromosomes plays a role in maintaining the expressed or silent state of some genes. The background for this hypothesis comes from fluorescence in situ hybridization studies of genomic DNA sequences performed in cells and tissues. Chromosome painting probes are available for all human chromosomes and for some chromosomes in many other mammalian species. In situ hybridization experiments were performed with such probes in tissue culture cells and revealed that there are distinct chromosomal domains for each of the chromosomes (Cremer et al. 1994; Sadoni et al. 1999; Strouboulis and Wolffe 1996). Although no specific position for each chromosome was found and no homologous pairing could be discerned, there appear to be distinct preferences for the nuclear boundary or the nuclear interior for specific chromosomes within the nuclei of dividing populations of human cells. Recently, it was shown that this level of nuclear architecture is altered in cells that have become either quiescent or senescent, suggesting that the spatial organization of the genome is plastic (Bridger et al. 2000; Croft et al. 1999). Such plasticity has already been demonstrated for specific gene loci, depending on their transcriptional or replication state (Brown et al. 1999; Dietzel et al. 1999; Kurz et al. 1996; Zirbel et al. 1993). These studies have been limited to mammalian tissue culture cells or circulating blood cells. It may be assumed that a much more distinct architecture is necessary in tissues and whole organisms where very specific developmentally controlled programs of gene expression are required.

We have studied the plasticity of chromosomes and specific genetic loci in developing Drosophila embryos and larvae and have shown that in this organism, there is very strong pairing of homologous chromosomes, which has specific consequences for gene expression. In addition, we determined that translocations of large chromosome segments did not inhibit this homologous association, showing that there is flexibility in both unpaired and paired chromosomes such that homologue searching occurs over a large volume (Gemkow et al. 1997; Gemkow et al. 1998). Other groups have also shown that genetic loci are repositioned within the nucleus of Drosophila cells when inserted in particular chromosomal contexts (Csink and Henikoff 1996; Dernburg et al. 1996), and that there is flexibility and dynamic repositioning throughout the cell cycle (Csink and Henikoff 1998). The dynamic repositioning of loci could be driven by the state of the histone and non-histone proteins that make up the chromatin. The new data presented in the lecture will address this hypothesis.

Bibliography for fluorescence in situ hybridization techniques:

Bickmore and Oghene 1996; Garciablanco et al. 1995; Gemkow 1999; Grünewald-Janho et al. 1996; Haaf and Ward 1995; Hendzel and Bazett-Jones 1997; Heng et al. 1997; Leger et al. 1994

Polycomb-Group Protein Repression Complexes, Cellular Memory

Early in Drosophila embryogenesis, both larval and adult body segment identities are determined by the expression patterns of the homeotic genes. The expression domains in the embryo are established by products of the segmentation genes, which decay by 5 to 7 hours of development. The spatial distribution of repression and expression of the homeotic genes, however, is maintained for up to 10 cell divisions before terminal cell differentiation. The class of genes responsible for the maintenance of repression is the Polycomb-group (PcG) (Paro 2000; Pirrotta 1998). Animals mutant for any of the about 15 members of the PcG identified thus far show multiple homeotic transformations resulting from the ectopic expression of homeobox genes outside their normally restricted domains. The maintenance of the determined state is a process required in every multicellular organism undergoing development (Akasaka et al. 2001). There is experimental evidence that the PcG proteins form a multi-protein chromatin-bound complex emanating from PRE (polycomb response element) sites in the genome in those cells in which the homeobox genes are repressed [see contributions at this meeting from Cavalli and Orlando; (Orlando et al. 1998; Strutt et al. 1997; Strutt and Paro 1997)].

We demonstrated in whole mount Drosophila embryos that PcG complexes are distributed throughout the interphase nuclei as >100 discreet loci and are excluded from the constitutive heterochromatin (Buchenau 1996; Buchenau et al. 1998). During embryogenesis, the PcG complexes disassociate from the chromatin at mitosis, leaving <2% of polyhomeotic protein, for example, attached to late metaphase and anaphase chromosomes. The reassociation of the various PcG proteins at telophase indicates that there is a hierarchy in the assembly of the PcG protein complexes. See Figure 1 [taken from (Buchenau et al. 1998)].

Figure 1. Distribution of PcG proteins across the cell cycle in developing Drosophila embryos. A gallery of single confocal sections showing the distribution of PC, PH and PSC during all distinguishable phases of the cell cycle for 2n nuclei. The images were extracted from whole mount embryos older than stage 8.

Figure 1

Distribution of PcG proteins across the cell cycle in developing Drosophila embryos. A gallery of single confocal sections showing the distribution of PC, PH and PSC during all distinguishable phases of the cell cycle for 2n nuclei. The images were extracted (more...)

The genes of the bithorax complex (BX-C) are differentially expressed during embryogenesis. We have developed a technique by which we can simultaneously label the PcG complexes as well as genes in the BX-C by FISH. We have recently reisolated a mutant Drosophila originally described by Bender, Ubx266 (Bender et al. 1985), where a rearrangement splits the BX-C such that the Ubx and Abd-B, abd-A genes reside on different chromosomes. This line is being used to study the colocalization of the complex and the genes during development.

The PcG complexes in imaginal discs (precursor nests of cells that will form the adult fly) show a distribution different from that in interphase embryos. In the discs, we find fewer and much larger complexes, suggesting that the individual repressed genes are forming into higher order complexes that may represent repression domains (see Figure 2). We have modified our microscope systems to accommodate whole embryos and larval tissues and have initiated FRAP and FCS experiments to compare the mobility of proteins in the complexes during different stages of Drosophila development.

Figure 2. Distribution of polyhomeotic protein, PH, in Drosophila 3rd instar larval imaginal disk nuclei from whole mount imaging. Extended focus image of 5 confocal sections at 0.5 μ intervals. Resolution in XY= 0.1 μ. Indirect immunofluorescence, Cy3 fluorophore.

Figure 2

Distribution of polyhomeotic protein, PH, in Drosophila 3rd instar larval imaginal disk nuclei from whole mount imaging. Extended focus image of 5 confocal sections at 0.5 μ intervals. Resolution in XY= 0.1 μ. Indirect immunofluorescence, (more...)

The Drosophila X Chromosome and the MSL Dosage Compensation Complex

The Drosophila genome has 120 Mbp, of which 22 Mbp reside in the X chromosome. It is estimated that there are 2,200 genes on this chromosome, of which some 800 would be essential. Females are XX and males X0. Dosage compensation ensures that males with a single X chromosome have the same amount of most X-linked gene products as females with two X chromosomes. In Drosophila, this equalization is achieved by a twofold enhancement of the level of transcription of the X in males relative to each X chromosome in females. The products of at least five genes, maleless (mle), male-specific lethal 1, 2, and 3 (msl-1, msl-2, msl-3), and males absent on the first (mof), are necessary for dosage compensation. The proteins produced by these genes form a complex that is preferentially associated with numerous sites on the X chromosome in somatic cells of males but not of females. RNA transcripts encoded by at least two different genes on the X chromosome (rox1 and rox2) function as the first entry sites for assembly of the MSL complex in males (Gu et al. 1998; Gu et al. 2000). It should be noted that although the MSL1 and MSL2 proteins are able to access the X chromosome at the entry sites and to recruit MLE (an ATPase), further complex assembly can only occur in the presence of the roX RNAs and the histone acetyltransferase activity of MOF. The spreading process may involve the acetylation of neighboring nucleosomes, thereby altering the conformation of adjacent chromatin and rendering it more accessible to the entry of additional MSL complexes. More recently, JIL-1, a novel chromosomal kinase that is up-regulated almost twofold on the male X chromosome in Drosophila, has been demonstrated to colocalize and physically interact with the msl dosage compensation complex proteins (Jin et al. 2000). Males must turn the male-specific lethal (msl)-mediated pathway of dosage compensation on, and females must keep it off (Chang and Kuroda 1998). Kuroda and coworkers demonstrated further that the MSL1, MSL2, and MSL3 proteins are associated in immunoprecipitations, chromatographic steps, and in the yeast two-hybrid system (Copps et al. 1998).

The MSL-complex can be visualized in developing male embryos and larvae by immunofluorescence using antibodies to the MSL proteins, GFP--MSL fusion proteins, and in situ hybridization to X chromosome sequences or to the roX transcripts (see Figure 3). Interestingly, we observe a gross rearrangment of the MSL protein complex in the nuclei of imaginal discs late in larval development. Instead of the domain structure that generally describes the X locus in embryogenesis and early larval development, the MSL complex is seen to move to the nuclear periphery and lie directly under the lamin-stained nuclear envelope (see Figure 4).

Figure 3. Distribution of MSL-1 in male Drosophila embryos. Extended focus stack of 12 sections at 0.5 μ intervals from a stage 8 embryo stained for lamin, red and MSL-1, green. Resolution in XY = 0.198 μ.

Figure 3

Distribution of MSL-1 in male Drosophila embryos. Extended focus stack of 12 sections at 0.5 μ intervals from a stage 8 embryo stained for lamin, red and MSL-1, green. Resolution in XY = 0.198 μ.

Figure 4. Disposition of MSL-1 in imaginal disc cells from 3rd instar male Drosophila larvae. Whole mount immunostaining and confocal imaging. Red, lamin; green, MSL-1 protein; blue, H2AvD-GFP chromatin.

Figure 4

Disposition of MSL-1 in imaginal disc cells from 3rd instar male Drosophila larvae. Whole mount immunostaining and confocal imaging. Red, lamin; green, MSL-1 protein; blue, H2AvD-GFP chromatin. A. early larval disc, extended focus of 4 confocal sections (more...)

Photophysical Methods to Extract Information about Molecular Proximity, Interactions, and Mobilities

FRAP: fluorescence recovery after photobleaching. FLIP: fluorescence loss in photobleaching. These methods use photobleaching of a chromophore to gain information about the mobility of molecules as well as their steady-state association/dissociation rates with complexes or compartments in living cells.

What FRAP/FLIP can do: 1) determine apparent diffusion rates of proteins in various cellular compartments; and 2) determine the percentage of a protein population that is freely diffusible.

What FRAP/FLIP cannot do: distinguish small changes in diffusion rates. For soluble proteins, the molecular mass must increase 8-fold to result in a 2-fold difference in the diffusion rate, i.e., one cannot detect dimerization by this method. For proteins in membranes, a 50-100-fold change in molecular weight is needed to change the diffusion rate 2-fold.

The following is a list of references to the methodology of FRAP/FLIP:

Gribbon et al. 1999; Gribbon and Hardingham 1998; Kaufman and Jain 1990; Kubitscheck et al. 1998; Periasamy and Verkman 1998; Peters and Kubitscheck 1999; Siggia et al. 2000; Storrie and Kreis 1996; Sullivan and Kay 1999; Wedekind et al. 1996

A small selection of some of the applications of these methods to problems in cell physiology and in particular, nuclear metabolism, follows:

Arrio-Dupont et al. 2000; Cardoso et al. 1999; Dundr et al. 2000; Hirschberg et al. 1998; Houtsmuller and Vermeulen 2001; Huang et al. 1998; Kruhlak et al. 2000; Misteli 2001; Misteli et al. 2000; Moir et al. 2000; Nehls et al. 2000; Partikian et al. 1998; Phair and Misteli 2000; Sullivan and Kay 1999; Sund and Axelrod 2000

FRET: fluorescence resonance energy transfer. This method allows the determination of molecular proximity. Imaging FRET is a very powerful method to prove complex formation and molecular associations within cells. It is limited to associations within ~150 Å and may be used as well to determine gene activity, ion concentrations, molecular integrity, or relative stoichiometry in complexes. Caveat! The absence of FRET does not necessarily mean that a complex does not exist but may simply reflect the fact that the two partners being probed are more than ~200 Å apart. FRET requires two molecules with chromophores that have spectral overlap in the emission spectrum of the donor and the excitation spectrum of the acceptor (note that the acceptor must not necessarily be fluorescent to make the measurement). FRET is measured by determining either 1) donor quenching, 2) acceptor sensitization, 3) decrease in the fluorescence lifetime of the donor, or 4) increase in the photobleaching time of the donor. Caveat!! The microscope and fluorophore systems must be calibrated, the undesired photon spillage into donor or acceptor channels quantitatively accounted for, and cells with "control" (non-interacting) proteins measured under the same conditions to extract proper quantitative measurements from the data.

The following is a list of references on the theory and practice of FRET measurements for microscopy:

Bastiaens and Jovin 1997; Clegg 1995; Clegg 1996; Clegg et al. 1992; Clegg et al. 1994; Gadella Jr. et al. 1994; Gadella et al. 1999; Hanley et al. 2001; Harpur et al. 2001; Jovin and Arndt-Jovin 1989; Miyawaki and Tsien 2000; Pepperkok et al. 1999; Selvin 2000; Zacharias et al. 2000

The use of fluorescent protein construct pairs for FRET has been reviewed recently by Miyawaki and Tsien (Miyawaki and Tsien 2000). In addition, Tsien's group has pioneered the use of chimeric molecules (cameleons) that monitor enzymatic activity or ionic or pH conditions in cellular compartments (Allen et al. 1999; Miyawaki et al. 1999; Miyawaki et al. 1997; Zlokarnik et al. 1998). One should be aware of the complex photophysical properties of GFP and its spectrally shifted mutants because changes in the emission spectrum may reflect non-FRET phenomena (Creemers et al. 2000; Creemers et al. 1999).

The following papers show the application of FRET to other cell biological problems:

Allen et al. 2000; Bastiaens and Jovin 1996; Bastiaens et al. 1996; Chamberlain and Hahn 2000; Gadella Jr. and Jovin 1995; Gadella et al. 1999; Goedhart et al. 1999; Li et al. 1999; Llopis et al. 1998; Ng et al. 1999; Raz et al. 1998; Zlokarnik et al. 1998

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