Information

2.0: DNA is Packaged into Chromatin - Biology

2.0: DNA is Packaged into Chromatin - Biology


We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

DNA can be highly compacted

If stretched to its full length, the DNA molecule of the largest human chromosome would be 85mm. Yet during mitosis and meiosis, this DNA molecule is compacted into a chromosome approximately 5µm long. Although this compaction makes it easier to transport DNA within a dividing cell, it also makes DNA less accessible for other cellular functions such as DNA synthesis and transcription. Thus, chromosomes vary in how tightly DNA is packaged, depending on the stage of the cell cycle and also depending on the level of gene activity required in any particular region of the chromosome.

Levels of compaction

There are several different levels of structural organization in eukaryotic chromosomes, with each successive level contributing to the further compaction of DNA (Figure (PageIndex{2})). For more loosely compacted DNA, only the first few levels of organization may apply. Each level involves a specific set of proteins that associate with the DNA to compact it. First, proteins called the core histones act as spool around which DNA is coiled twice to form a structure called the nucleosome. Nucleosomes are formed at regular intervals along the DNA strand, giving the molecule the appearance of “beads on a string”. At the next level of organization, histone H1 helps to compact the DNA strand and its nucleosomes into a 30nm fibre. Subsequent levels of organization involve the addition of scaffold proteins that wind the 30nm fibre into coils, which are in turn wound around other scaffold proteins.

Chromatin Packaging Varies inside the Nucleus: Euchromatin & Heterochromatin

Chromosomes stain with some types of dyes, which is how they got their name (chromosome means “colored body”). Certain dyes stain some regions along a chromosome more intensely than others, giving some chromosomes a banded appearance. The material that makes up chromosomes, which we now know to be proteins and DNA, is called chromatin. Classically, there are two major types of chromatin, but these are more the ends of a continous and varied spectrum. Euchromatin is more loosely packed, and tends to contain genes that are being transcribed, when compared to the more densely compacted heterochromatin, which is rich in repetitive sequences and tends not to be transcribed. Heterochromatin sequences include short, highly-repetitive sequences called satellite DNA, which acquired their name because their bouyant density is distictly different from the main band of DNA following ultracentrifugation.

Morphological features of Chromosomes

Chromosomes also contain other distinctive features such as centromeres and telomeres. Both of these are usually heterochromatin. In most cases, each chromosome contains one centromere. These sequences are bound by centromeric proteins that link the centromere to microtubules that transport chromosomes during cell division. Under the microscope, centromeres of metaphase chromosomes can sometimes appear as constrictions in the body of the chromosome (Figure (PageIndex{3})) and are called primary (1°) constrictions. If a centromere is located near the middle of a chromosome, it is said to be metacentric, while an acrocentric centromere is closer to one end of a chromosome, and a telocentric chromsome is at, or near, the very end. In contrast, some species have a holocentric centromere, where no single centromere can be defined and the entire chromsome acts as the centromere. Telomeres are repetitive sequences near the ends of linear chromosomes, and are important in maintaining the length of the chromosomes during replication, and protecting the ends of the chromosomes from alterations.

It is essential to describe the similarity between chromosomes using appropriate terminology (Fig 2.4). Homologous chromosomes are typically pairs of similar, but non-identical, chromosomes in which one member of the pair comes from the male parent, and the other comes from the female parent. Homologs contain the same gene loci but not necessarily the same alleles. Non-homologous chromosomes contain different gene loci, and are usually distinguishable based on cytological features such as length, centromere position, and banding patterns.

An unreplicated chromosomes can undergo replication, to produce a replicated chromosome that has two sister chromatids, which are physically connected to each other at the centromere and remain joined until cell division. Because a pair of sister chromatids is produced by the replication of a single DNA molecule, their sequences are essentially identical (same alleles), differing only because of DNA replication errors. On the other hand, non-sister chromatids come from two separate, but homologous chromosomes, and therefore usually contain the same gene loci in the same order, but do not necessarily have identical DNA sequences (allelic differences).

The decondensed chromosomes are not randomly arranged in within the interphase nucleus. They often have specific locations within the nucleus and relative to one another (Figure (PageIndex{5}))

Chromosome and DNA Replication

When the cell enters S-phase in the cell cycle (G1-S-G2-M) all the chromosomal DNA is replicated. This is done by enzymes called DNA polymerases. All DNA polymerases synthesize new strands by adding nucleotides to the 3'OH group present on the previous nucleotide. For this reason they are said to work in a 5' to 3' direction. DNA polymerases use a single strand of DNA as a template upon which it will synthesize the complementary sequence. This works fine for the middle of chromosomes - DNA-directed DNA polymerases travel along the original DNA strands making complementary strands (Figure (PageIndex{6})a).

DNA replication in both prokaryotes and eukaryotes begins at an Origin of Replication (Ori). Origins are specific sequences on specific positions on the chromosome. In E. coli, the OriC origin is ~245 bp in size. Chromosome replication begins with the binding of the DnaA initiator protein to an AT-rich 9-mer in OriC and melts the two strands. Then DnaC loader protein helps DnaB helicase protein extend the single stranded regions such that the DnaG primase can initiate the synthesis of an RNA primer, from which the DNA polymerases can begin DNA synthesis at the two replication forks. The forks continue in opposite directions until they meet another fork or the end of the chromosome (Figure (PageIndex{7})).

The ends of linear chromosomes present a problem – at each end one strand cannot be completely replicated because there is no primer to extend. Although the loss of such a small sequence would not be a problem, the continued rounds of replication would result in the continued loss of sequence from the chromosome end to a point were it would begin to loose essential gene sequences. Thus, this DNA must be replicated. Most eukaryotes solve the problem of synthesizing this unreplicated DNA with a specialized DNA polymerase called telomerase, in combination with a regular polymerase. Telomerases are RNA-directed DNA polymerases. They are a riboprotein, as they are composed of both protein and RNA. As Figure (PageIndex{6})b shows, these enzymes contain a small piece of RNA that serves as a portable and reusable template from which the complementary DNA is synthesized. The RNA in human telomerases uses the sequence 3-AAUCCC-5' as the template, and thus our telomere DNA has the complementary sequence 5'-TTAGGG-3' repeated over and over 1000’s of times. After the telomerase has made the first strand a primase synthesizes an RNA primer and a regular DNA polymerase can then make a complementary strand so that the telomere DNA will ultimately be double stranded to the original length (Figure (PageIndex{8})). Note: the number of repeats, and thus the size of the telomere, is not set. It fluctuates after each round of the cell cycle. Because there are many repeats at the end, this fluctuation maintains a length buffer – sometimes it’s longer, sometimes it’s shorter – but the average length will be maintained over the generations of cell replication.

In the absence of telomerase, as is the case in human somatic cells, repeated cell division leads to the “Hayflick limit”, where the telomeres shorten to a critical limit and then the cells enter a senescence phase of non-growth. The activation of telomerase expression permits a cell and its descendants to become immortal and bypass the Hayflick limit. This happens in cancer cells, which can form tumours as well as in cells in culture, such as HeLa cells, which can be propagated essentially indefinitely. HeLa cells have been kept in culture since 1951.

Eukaryote Chromosomes have Multiple Origins of Replication

In prokaryotes, with a small, simple, circular chromosome, only one origin of replication is needed to replicate the whole genome. For example, E. coli has a ~4.5 Mb genome (chromosome) that can be duplicated in ~40 minutes assuming a single origin, bi-directional replication, and a speed of ~1000 bases/second/fork for the polymerase.

However, in larger, more complicated eukaryotes, with multiple linear chromosomes, more than one origin of replication is required per chromosome to duplicate the whole chromosome set in the 8-hours of S-phase of the cell cycle. For example, the human diploid genome has 46 chromosomes (6 x 109 basepairs). The shortest chromosomes are ~50 Mbp long and so could not possibly be replicated from one origin. Additionally, the rate of replication fork movement is slower, only ~100 base/second. Thus, eukaryotes contain multiple origins of replication distributed over the length of each chromosome to enable the duplication of each chromosome within the observed time of S-phase (Fig 2.9).


In eukaryotic cells, DNA is packaged into a complex macromolecular structure called chromatin. Wang et al. have developed an imaging method to map the position of multiple regions on individual chromosomes, and the results confirm that chromatin is organized into large contact domains called TADS (topologically associating domains). Unexpectedly, though, folding deviates from the classical fractal-globule model at large length scales.

The spatial organization of chromatin critically affects genome function. Recent chromosome-conformation-capture studies have revealed topologically associating domains (TADs) as a conserved feature of chromatin organization, but how TADs are spatially organized in individual chromosomes remains unknown. Here, we developed an imaging method for mapping the spatial positions of numerous genomic regions along individual chromosomes and traced the positions of TADs in human interphase autosomes and X chromosomes. We observed that chromosome folding deviates from the ideal fractal-globule model at large length scales and that TADs are largely organized into two compartments spatially arranged in a polarized manner in individual chromosomes. Active and inactive X chromosomes adopt different folding and compartmentalization configurations. These results suggest that the spatial organization of chromatin domains can change in response to regulation.

The spatial organization of chromatin, such as chromatin domains, chromatin loops, associations of chromatin with nuclear structures, and chromosome territories, plays an important role in essential genome functions (16). However, many gaps remain in our understanding of the three-dimensional (3D) folding of individual chromosomes in the nucleus. Recently, chromosome-conformation-capture methods such as Hi-C (4, 7) have revealed a wealth of structural insights for interphase chromosomes. For example, chromatin is organized into topologically associating domains (TADs) or contact domains that are hundreds of kilobases (kb) in size (811). These domains tend to spatially segregate from each other (9, 12, 13) and, in Drosophila, correspond to the banding patterns of polytene chromosomes (14, 15). At length scales from several hundred kilobases to several megabases, the power-law scaling of Hi-C contact frequency is consistent with a fractal-globule polymer model (7, 16), whereas, within TADs, Hi-C contact maps are better described by a loop-extrusion model (17, 18). Superresolution imaging shows that chromatin domains in different epigenetic states adopt distinct folding configurations with different power-law scaling properties (13). Whether the ideal fractal-globule model can describe chromatin at length scales beyond several megabases remains an open question (19, 20). Hi-C analyses have also revealed two multi-TAD compartments, compartments A and B, that are enriched with active and inactive chromatin, respectively (7, 21). However, because the contact maps used to identify these compartments were derived from ensemble averaging of many chromosomes, it is unclear whether the A and B compartments are structures that exist in individual chromosomes inside single cells and, if so, how the two compartments are spatially arranged with respect to each other. To answer questions regarding the spatial organization of individual chromosomes requires methods that directly visualize the conformation of single chromosomes in single cells.

Fluorescence in situ hybridization (FISH) provides a powerful means to directly image the spatial organization of chromosomes, especially when used to simultaneously target two or more genomic loci (e.g., 19, 20, 2224). In one effort, a three-color barcoding approach has been used to simultaneously label multiple chromatin loci to trace the conformation of a chromosome arm in Drosophila blastoderm embryos (24). Nonetheless, routine tracing of the complex 3D folding path of chromosomes has remained challenging because of the difficulties associated with simultaneously imaging and unambiguously identifying many genomic regions on interphase chromosomes. Here, we report a multiplexed FISH method that enables sequential imaging of many genomic regions for 3D tracing of individual chromosomes in the nucleus and the use of this method to study the spatial arrangements of TADs and compartments in chromosomes 20, 21, 22, and X of human diploid (XX) IMR90 cells.

To map the 3D spatial positions of TADs, delineated here as genomic domains based on ensemble-average Hi-C maps (8), along an entire chromosome, we labeled the central 100-kb regions of TADs using a dual-oligonucleotide version of Oligopaints (25, 26), wherein each TAD was targeted with 1000 distinct “primary” oligonucleotide probes and a companion “secondary” probe (Fig. 1A and fig. S1). Each of the 1000 primary probes consisted of a unique targeting sequence complementary to a given sequence within the TAD and a nongenomic region, called Mainstreet, that contained a readout sequence shared by all 1000 probes but unique for each TAD. The secondary probe contained a sequence complementary to the readout sequence on Mainstreet. We used the genomic coordinates of TADs derived from Hi-C (8) to design the targeting sequences of primary probes and produced these probes with a high-yield enzymatic amplification method (27). We exploited a similar hybridization and imaging protocol as we previously described in multiplexed error-robust fluorescence in situ hybridization (MERFISH) (27) but with some modifications. First, we hybridized all primary probes to the chromosome of interest (fig. S1, Hyb 0), imaged the sample, and located the chromosome in the nucleus. We then photobleached the sample and performed a series of secondary hybridizations, separated by photobleaching, to sequentially label and image individual TADs (Fig. 1A and fig. S1, Hyb 1, Hyb 2, and so on). Each round of secondary hybridization employed two different secondary probes, respectively labeled with two spectrally distinct dyes, enabling us to visualize two TADs simultaneously using two-color, 3D fluorescence imaging with z-stepping. The centroid positions of the 3D images of individual TADs were used to approximate their positions in x, y, and z.

(A) A simplified scheme of the imaging approach. All primary probes are first hybridized to the targeted chromosome, after which secondary probes targeting each TAD are sequentially hybridized to the readout sequences on the primary probes, imaged, and then bleached. In each round of secondary hybridization, two different secondary probes tagged with dyes of different colors allowed simultaneous visualization of two TADs. More details are depicted in fig. S1. (B) Image of an IMR90 cell after the primary hybridization (Hyb 0) with primary probes targeting all TADs in Chr21. The two bright patches, one marked by a yellow box, correspond to the two copies of Chr21 in this diploid cell. (C) Images of the yellow-boxed region in (B) after each round of secondary hybridization (Hyb 1 to 17). (D) Positions of the 34 TADs of the chromosome were plotted as red dots overlaid on the Hyb 0 image. Scale bars in (B) to (D), 2 μm. (E) TAD positions plotted in 3D. (F) Mean spatial distance matrix for the 34 TADs, with each element of the matrix corresponding to the mean spatial distance between a pair of TADs. (G) Inverse Hi-C contact frequency between each pair of TADs versus their mean spatial distance. The correlation coefficient (R) and the slope of a fitted line (k) are shown. Contact frequency is calculated as the total Hi-C counts between two TADs normalized to their genomic lengths (8). (H) Mean spatial distance versus genomic distance for all pairs of TADs. The lines are power-law function fits with either a predefined scaling exponent (S = 1/3, green) or with S as a fitting parameter (red). Data from 120 individual chromosomes were used to generate (F) to (H).

We first used this method to image chromosome 21 (Chr21) in interphase IMR90 cells. The Hyb 0 image showed that, when imaged together, the fluorescent signals from all 34 TADs of Chr21 coalesced into a continuous patch (Fig. 1B). The 17 rounds of secondary hybridization then allowed us to image each of the 34 TADs separately (Fig. 1C), determine the 3D position of each TAD, and trace the 3D path of this chromosome at the TAD level (Fig. 1, D and E). To characterize the organization of Chr21, we traced 120 copies of Chr21 in many cells, calculated the mean spatial distance between each pair of TADs (averaged over all 120 chromosomes), and constructed a pair-wise mean spatial distance matrix for the 34 TADs (Fig. 1F).

To compare our measurements with previous Hi-C data (8), we correlated the mean spatial distance matrix with the corresponding Hi-C contact frequency matrix of Chr21 (fig. S2). Notably, the mean spatial distance showed high correlation with the inverse contact frequency between TADs, with a Pearson correlation coefficient of 0.91 across nearly three orders of magnitude in contact frequency (Fig. 1G). Such a strong correlation between the results from two different methods provided a cross validation for both methods at the TAD-to-chromosome length scales probed in this work. The relationship between the spatial distance and contact frequency also provides a valuable measure for exploring chromosome organization. A mean-field approximation predicts that the contact frequency should be inversely proportional to the third power of the mean spatial distance, whereas the power for real chromatin is expected to be bigger than 3 (19). Our data showed that the Hi-C contact frequency was inversely proportional to the fourth power of the mean spatial distance [Fig. 1G, scaling exponent k = 4.1 ± 0.1, 95% confidence interval (CI), N = 120 chromosomes]. We also analyzed the distributions of the spatial distances between pairs of TADs (fig. S3) and found that the Hi-C contact frequency scaled linearly with the probability of two TADs coming into spatial proximity (fig. S4). These results suggest a calibration function to convert Hi-C contact frequencies into mean spatial distances at TAD-to-chromosome length scales, although it remains to be determined whether this calibration extends to sub-TAD scales where the correlation between the Hi-C contact frequency and spatial proximity may be weaker (28).

In addition, our data showed that the mean spatial distance between TADs scaled with their genomic distance to about one-fifth power (Fig. 1H, scaling exponent S = 0.21 ± 0.01, 95% CI, N = 120 chromosomes). This value deviated from the one-third power expected from the ideal fractal-globule polymer model (19). The deviation was most pronounced for large genomic distances, whereas data points with genomic distances less than 7 Mb showed a scaling exponent close to one-third (fig. S5), consistent with previous results (7, 19). Interestingly, a previous simulation of confined, unknotted, finite-sized polymers showed a deviation of the scaling exponent from one-third at large length scales (29), suggesting a possible physical model to explain our experimental observation.

Next, we determined whether the spatial positions of TADs are partitioned into distinct compartments by implementing a normalization analysis similar to that performed for the Hi-C data (7). First, we normalized the mean spatial distance matrix to the expected spatial distance at each genomic distance as predicted by the power-law scaling shown in Fig. 1H. The normalized spatial distance matrix showed a pattern with alternating regions of large and small values (Fig. 2A), suggesting the existence of two subgroups of TADs. Next, we calculated the Pearson correlation coefficient between each pair of columns in the normalized distance matrix, defined this coefficient as the correlation between the two corresponding TADs, and constructed a Pearson correlation matrix for all TAD pairs (Fig. 2B). This Pearson correlation matrix showed a plaid pattern, consistent with the existence of two compartments with TADs from the same compartment being positively correlated. For comparison, we used a similar approach (7) to analyze the Hi-C data (8) and obtained a nearly identical Pearson correlation matrix (Fig. 2C), suggesting that the two compartments observed in our imaging data correspond to the A and B compartments identified by Hi-C analysis (7, 21). To assign each TAD to a compartment, we performed a principal components analysis on the Pearson correlation matrix derived from the normalized spatial distances and assigned TADs with positive and negative values along the first principal component to compartments A and B, respectively (Fig. 2D). Nearly identical assignment was obtained by applying the principal components analysis directly to the normalized spatial distance matrix (fig. S6). We further observed that histone modifications for active chromatin (30, 31) and inactive chromatin (32) were enriched in compartments A and B, respectively (fig. S7), consistent with previous Hi-C analysis (7). We then analyzed the scaling relationship between inverse Hi-C contact frequency and mean spatial distance for pairs of TADs that are either within the same compartment or cross-compartment and found that cross-compartment TAD pairs gave a moderately higher scaling exponent (fig. S8).

(A) Normalized spatial distance matrix for the 34 TADs, normalized over the expected spatial distances determined by the power-law function fit in Fig. 1H (red line). (B) Pearson correlation matrix of the 34 TADs, determined from the normalized spatial distance matrix in (A). (C) Pearson correlation matrix of the 34 TADs calculated from previous Hi-C data (8). (D) Assignment of TADs to compartment A (red bars) or compartment B (blue bars) based on a principal components analysis of the Pearson correlation matrix shown in (B). (E) (Left panels) Spatial position maps of compartment-A TADs (red) and compartment-B TADs (blue) in two individual chromosomes. For better visualization, the chromosomes were rotated so that the polarization axis connecting the centroids of compartments A and B is aligned along the z axis. (Right panels) Corresponding 3D convex hull plots. (F) Polarization index values measured for individual chromosomes (observed) in comparison with those derived from a randomization control where the compartment assignments were randomized while maintaining the total number of TADs in each compartment. The nonzero control values arose from fluctuations associated with the finite number of TADs per chromosome, which provides a baseline for comparison. Each dot corresponds to the polarization index of a single chromosome, the red lines represent the median values, and the blue boxes represent the 25% to 75% quantiles. **P < 0.001 (Wilcoxon test). Data from 120 individual chromosomes were used to generate (A), (B), (D), and (F).

The above population-averaged analyses cannot reveal whether the higher correlation observed between TADs in the same compartment represents transient proximity between these TADs or whether the two compartments are physical structures that exist in individual chromosomes, nor can they reveal how compartments are spatially arranged—e.g., whether one compartment wraps around the other to form a radial organization within a single chromosome or whether the two compartments are arranged in a side-by-side, polarized manner. To address these questions, we examined the spatial positions of the central regions of TADs in single chromosomes. Notably, most individual chromosomes in single cells showed a spatially polarized arrangement of compartment-A and compartment-B TADs (Fig. 2E). To quantify the polarized separation of the compartments in individual chromosomes, we defined a polarization index as , where VA and VB are the convex hull volumes of the two compartments and VS is their shared volume. If the two compartments perfectly overlap with each other, or if one compartment wraps around the other, the polarization index should equal zero on the other hand, if the two compartments are completely separated in space in a polarized manner, the polarization index should equal 1 (fig. S9). The measured polarization index values of Chr21 were indeed close to 1, with a median value of 0.86, substantially larger than the values derived from a randomization control (Fig. 2F).

To investigate whether the above findings were chromosome-specific, we traced the positions of the central 100-kb regions of TADs in Chr22 and Chr20 by imaging all 27 TADs in Chr22 and 30 of the 60 TADs (every other one) in Chr20 and found conclusions similar to those described for Chr21. First, the Hi-C contact frequency was inversely proportional to the fourth power of the mean spatial distance between TADs (figs. S10A and S11A). Second, the mean spatial distance between TADs scaled with genomic distance to a similar, albeit slightly smaller, power than in Chr21 (Fig. 3, A and B), substantially deviating from the one-third power predicted by the ideal fractal-globule model. Third, analysis based on spatial distances showed that TADs in Chr22 and Chr20 were partitioned into two spatial compartments (Fig. 3, C and D, and figs. S10, B to E, and S11, B to E), with assignments nearly identical to those obtained from our analysis on Hi-C data. These two compartments were again spatially organized in a polarized, side-by-side manner in individual chromosomes (Fig. 3, E to H), although the degree of polarized separation is moderately smaller in Chr20. Whether these findings extend to all other autosomes remains to be determined.

(A and B) Mean spatial distance versus genomic distance for Chr22 (A) and Chr20 (B). Power-law function fits are shown as red lines, and the scaling exponents (S) are shown. (C and D) Compartment assignments of TADs based on principal components analyses of the Pearson correlation matrix for Chr22 (fig. S10D) and Chr20 (fig. S11D). Blue bars, compartment B. Red bars, compartment A. (E and F) Spatial position maps of compartment-A TADs (red) and compartment-B TADs (blue) in single chromosomes for Chr22 (E) and Chr20 (F), plotted without (left) or with (right) 3D convex hulls. (G and H) Polarization index values measured for individual chromosomes for Chr22 (G) and Chr20 (H) (observed) in comparison with those of the randomization control (control). The dots, red lines, and blue boxes are defined as in Fig. 2F. **P < 0.001 (Wilcoxon test). Data from

150 individual chromosomes were used to generate (A), (C), and (G), and data from

110 individual chromosomes were used to generate (B), (D), and (H).

Finally, we traced the positions of the central 100-kb regions of TADs in the X chromosome (ChrX). We imaged 40 TADs (out of 86 total) spanning the whole chromosome at relatively uniform intervals. It is known that one of the two copies of ChrX in female mammalian cells undergoes X-inactivation (33, 34). We used TAD coordinates obtained from the combined Hi-C data (8) of both active and inactive copies of ChrX (Xa and Xi) to determine labeling sites but note that the TAD structures are attenuated or absent on Xi (9, 35). We distinguished Xa and Xi by immunostaining of macroH2A.1 (fig. S12), a histone variant enriched in Xi (35). The mean spatial distance matrices of Xi and Xa were strikingly different, with the Xi matrix elements being substantially more homogeneous and mostly having smaller values than the Xa matrix elements (fig. S13, A and B). Indeed, fitting a power-law function to the spatial versus genomic distance plot yielded a very small scaling exponent of S = 0.074 ± 0.003 (95% CI, N = 95 chromosomes) for Xi (Fig. 4A), whereas the scaling exponent for Xa (S = 0.22 ± 0.01, 95% CI, N = 95 chromosomes) remained similar to those of Chr20, Chr21, and Chr22 (Fig. 4B). These observations suggest that Xi not only was more compact (36) but also adopted a spatially more intermixed chromatin arrangement with more homogeneous interloci distances, reminiscent of the chromatin organization observed for Polycomb-repressed domains using superresolution imaging (13). Given the enrichment of Polycomb group proteins on Xi (33, 34), these observations suggest a potentially general mechanism to induce such a compact and highly intermixed chromatin folding configuration.

The TAD structures are attenuated or absent on Xi (9, 35) and hence, for Xi, the term “TAD” simply represents imaged genomic loci. (A and B) Mean spatial distance versus genomic distance for Xi (A) and Xa (B). Power-law function fits are shown as red lines, and the scaling exponents (S) are shown. (C and D) Compartment assignments for Xi (C) and Xa (D), based on principal components analyses of the Pearson correlation matrix for Xi (fig. S13C) and Xa (fig. S13D). Positions of the DXZ4 macrosatellite in Xi and the p and q arms in Xa are indicated. (E and F) Spatial position maps of TADs in single Xi (E) and Xa (F) chromosomes, without (left) or with (right) 3D convex hulls. (G and H) Polarization index measured for Xi (G) and Xa (H) (observed) in comparison with those of the randomization control (control). The dots, red lines, and blue boxes are defined as in Fig. 2F. **P < 0.001 (Wilcoxon test). Data from 95 individual chromosomes were used to generate (A) to (D), (G), and (H).

Notably, ChrX also formed two compartments, but the compartmentalization schemes were different for Xi and Xa. Consistent with previous allele-specific Hi-C analyses (21, 35, 37), Xi was largely partitioned into two contiguous compartments (also called superdomains or megadomains) separated on the genomic map by the DXZ4 macrosatellite (Fig. 4C and fig. S13, C, E, and G). Such a scheme might result from the ability of the DXZ4 element to recruit the chromatin insulator CTCF to Xi but not to Xa (38). The Xa TADs were also partitioned into two spatial compartments, but the two compartments corresponded instead to the p and q arms of the chromosome (Fig. 4D and fig. S13, D, F, and H). Interestingly, the two compartments in both Xa and Xi were again spatially organized in a polarized, side-by-side manner in individual chromosomes (Fig. 4, E to H). However, the degree of polarized segregation was notably smaller for Xi (Fig. 4G), consistent with our observation of more intermixed chromatin in Xi. It is worth noting that within the individual arms of Xa, TADs were further partitioned into two subcompartments (fig. S14, A to C), one of which appeared to be relatively enriched with histone modifications for active chromatin (fig. S14D), implying that these subcompartments potentially correspond to the A and B compartments.


What is chromatin?

The DNA of prokaryotic cells posses a minimal amount of information, so it is simply distributed in a circular form over the cytoplasm. However, the DNA of eukaryotes contains millions of pieces of hereditary information. Therefore, it&rsquos important to organize them properly in order to fit into the nucleus. Chromatin is a way to organize the genetic information to form the blueprint of life. It helps to pack the DNA into a small volume, so that it resides within the nucleus, with all the genetic information contained safely. It prevents the DNA from becoming tangled and plays a major role in reinforcing the DNA during cell division by regulating gene expression, facilitating DNA replication and preventing damage.

Chromatin strand (Photo Credit : Juan Gaertner/ Shutterstock)


Tsukiyama Lab

Chromatin regulation for cell cycle and cell division control
We investigate how chromatin structure is regulated in vivo. In eukaryotic cells, DNA is packaged into chromatin. This allows compact storage of the genome, but limits the access of DNA binding proteins to their targets. Therefore, chromatin structure strongly influences all processes that rely on protein-DNA interactions, including transcription, DNA replication, repair and recombination, and mis-regulation of chromatin structure can lead to diseases such as cancer. One of the major challenges in studying chromatin regulation is to elucidate how chromatin regulation affects such a wide variety of processes in biological contexts, such as cell cycle control and cell differentiation. We are particularly interested in understanding mechanisms of chromatin regulation within these important biological contexts. We use a diverse set of approaches, including genomics, molecular genetics, cell biology and biochemistry. Graduate students in the lab learn how to perform a wide variety of techniques, including deep sequencing and bioinformatic analyses.


Packaging and unpacking of the genome

DNA represents a dynamic form of information, balancing efficient storage and access requirements. Packaging approximately 1.8m of DNA into something as small as a cell nucleus is no mean feat, but unpacking it again to access the required sections and genes? That requires organisation.

In a nutshell, this is achieved through DNA condensed and packaged as chromatin, a complex of DNA and proteins called histones, which is constantly modified as the DNA is accessed. The histone proteins need constant replacement to maintain the correct chromatin structure required for all DNA related processes in the cell.

To understand more about the importance of histone replacement, researchers at the Babraham Institute and MRC Clinical Sciences Centre used developing mouse egg cells, oocytes. Developing oocytes provide a system where the mechanics of how DNA is packaged into cells can be explored in the absence of DNA replication, as egg cells do not divide. However, their genomes are highly active as the development of the egg involves widespread turning on and off of genes and DNA modification before the mature egg cell is ready for fertilisation. The work, published in the latest issue of Molecular Cell, relied on the Institute's expertise in single cell analysis, allowing accurate mapping of the epigenetic landscape in precious cells.

The researchers deleted a histone chaperone protein -- one of a group of proteins that are responsible for replacing histones in the chromatin structure -- and analysed the effects on egg cell development, DNA integrity and accumulation of DNA methylation.

"Oocytes lacking the Hira histone chaperone showed severe developmental defects which often led to cell death." said Dr Gavin Kelsey, research group leader in the Institute's Epigenetics programme and author on the paper. "The whole system is disrupted, eggs accumulate DNA damage and the altered chromatin means that genes cannot be efficiently silenced or activated. But we also uncovered an intricate relationship between the different epigenetic systems operating in the oocyte, where failure to ensure normal histone levels severely compromised deposition of methylation on the underlying DNA."

The research addresses the importance of histone turnover in maintaining genomic fidelity and adds to our understanding about the mechanisms in place to protect the integrity of the genome as it is remodelled and reshaped. Studying this in the context of the developing oocytes provides new insights into our dynamic genome, unclouded by the complications of DNA replication, and also reveals how important maintaining chromatin dynamics is to the integrity of our gametes.


Department of Cancer Biology

RESEARCH INTERESTS:

In eukaryotes, DNA is packaged into chromatin, the basic unit of which is the nucleosome. Chromatin structure plays a critical role in the regulation of gene expression by imposing topological constraints and by creating a barrier for transcriptional regulators. SWI/SNF enzymes are multiprotein complexes that alter chromatin structure in an ATP dependent manner and are involved in the regulation of gene expression. Components of the SWI/SNF complex are essential for mouse development and play important roles in several human cancers.

Cellular differentiation is characterized by the activation of previously silent genes embedded in repressive chromatin structure that is inaccessible to the transcriptional machinery. SWI/SNF enzymes interact with master regulators of differentiation to disrupt chromatin structure and activate lineage specific gene expression. We have previously determined that SWI/SNF enzymes interact with the master regulator of melanocyte differentiation, Microphthalmia Transcription Factor (MITF). MITF regulates the expression of genes that encode the enzymes needed for melanin synthesis, melanosome structure, and melanocyte survival. The main focus of my research is to study the functional role of SWI/SNF enzymes in the regulation of gene expression during melanocyte differentiation and to determine how SWI/SNF function is de-regulated in melanoma.

Member of the mentoring faculty for the Biomedical Sciences Graduate Program (Cancer Biology Track).

EDUCATION:

Ph.D. 1998 University of California, Davis, Davis, CA
B.S. 1984 Cornell University, Ithaca, NY

RECENT ACADEMIC APPOINTMENTS:

2012-pres Associate Professor, Cancer Biology, University of Toledo Health Scicence Campus
2005-2012 Assistant Professor, Biochemistry & Cancer Biology, University of Toledo Health Science Campus
2004-2005 Research Assistant Professor, Department of Cell Biology, University of Massachusetts Medical School
1998-2004 Postdoctoral Fellow, Department of Cell Biology, University of Massachusetts Medical School,
1990-1997 Research & Teaching Assistant, Department of Plant Pathology, Univesity of California, Davis
1988-1990 Research & Teaching Assistant, Department of Biological Sciences, California State University, Fullerton


First 3-D atomic view of key genetic processes

In a landmark study to be published in the journal Nature, scientists have been able to create the first picture of genetic processes that happen inside every cell of our bodies. Using a 3-D visualization method called X-ray crystallography, Song Tan, an associate professor of biochemistry and molecular biology at Penn State University, has built the first-ever image of a protein interacting with the nucleosome -- DNA packed tightly into space-saving bundles organized around a protein core. The research is expected to aid future investigations into diseases such as cancer.

As the genetic blueprint of life, DNA must be deciphered or "read," even when densely packed into nucleosomes. The nucleosome is therefore a key target of genetic processes in a cell and a focus of scientific investigations into how normal and diseased cells work. Previous studies at Penn State and other research institutions led to the discovery of chromatin enzymes -- proteins that act to turn specific genes on or off by binding to the nucleosome. Since the three-dimensional structure of the nucleosome was determined 13 years ago, scientists have wondered how chromatin enzymes recognize and act on the nucleosome to regulate gene expression and other processes in a cell. "We needed to visualize how these enzymes are able to read such a complicated structure as the nucleosome," Tan said.

To tackle this problem, Ravindra D. Makde, a postdoctoral member of the research team led by Tan, grew molecular crystals of the protein RCC1 (regulator of chromosome condensation, a protein critical for proper separation of chromosomes during cell division) bound to the nucleosome, and used X-ray crystallography to determine the atomic structure of the complex. "Our results showed that the RCC1 protein binds to opposite sides of the nucleosome -- similar to pedals positioned on a tricycle wheel." The structure provides atomic details of how an enzyme can recognize both DNA and components of the protein core of the nucleosome. Unexpectedly, the structure also showed how DNA can stretch as it wraps into a nucleosome. "These findings provide the basis for understanding how RCC1 and other chromatin enzymes interact with DNA as it is packaged into chromatin in our cells," Tan said.

The investigations were performed at the Penn State Center for Eukaryotic Gene Regulation, a multidisciplinary center focused on understanding the molecular basis for how genes are turned off and on in our bodies. "For years, the research community has been at an impasse," said Frank Pugh, Director of the center and the Willaman Professor in Molecular Biology at Penn State. "We were limited to only speculating how cellular proteins might bind the nucleosome. Now, with this structure, we are one step closer to understanding how cells read chromatin to regulate gene expression."

After nearly a decade of working to this goal, Tan and his team are excited to see the intricate interactions between a chromatin protein and the nucleosome. They are, however, even more enthusiastic about future prospects. "Our goal now is to determine the structures of other biologically and medically important chromatin enzymes bound to the nucleosome," said Tan. "We anticipate such studies will explain fundamental genetic processes and provide the basis for new therapeutics against human diseases such as cancer."

In addition to Tan and Makde, other researchers who contributed to this project include Joseph R. England, a Penn State undergraduate when he started this research and currently an MD/Ph.D. student at Temple University, and Hemant P. Yennawar, a senior research associate in the Department of Biochemistry and Molecular Biology at Penn State. This research was funded, in part, by the National Institutes of Health.

Story Source:

Materials provided by Penn State. Note: Content may be edited for style and length.


How to study epigenetics

​Epigenetic regulation occurs on many interacting levels, and it is essential to examine all of these levels in parallel to understand epigenetics contributions to biological processes. Tackling epigenetic studies from multiple angles with redundancy is key to ensuring accurate results.

Here we focus on five essential aspects of epigenetic regulation.

2. Histone modifications
Histones are proteins responsible for packaging DNA. A variety of mechanisms and modify histones, including acetylation, methylation, and phosphorylation, to control their interactions with DNA and therefore DNA structure and gene activation. Examining histone modifications, and the activity of enzymes that control these modifications can reveal mechanisms of epigenetic regulation and dysregulation at specific gene sites or across the genome at large​

3. ChIP guide
​Many different types of proteins bind to DNA to either directly or indirectly to regulate chromatin conformation and gene transcription. Identifying the presence or absence of such proteins in specific regions or across the genome can provide help build a complete picture of epigenetic regulation and dysregulation, as well as point to particular players and pathways involved. We can study these aspects of epigenetic regulation with chromatin immunoprecipitation (ChIP).​

4. Chromatin profiling using CUT&RUN and CUT&Tag
The Henikoff lab has recently developed two new chromatin profiling methods: Cleavage Under Targets and Release Using Nuclease (CUT&RUN) and Cleavage Under Targets and Tagmentation (CUT&Tag) which provide an exciting advance because they overcome many of the drawbacks of conventional and widely used chromatin immunoprecipitation (ChIP) methods.

5. DNA modifications
​​Throughout the DNA sequence, many chemical modifications exist. The most well-studied of these is 5-methylcytosine (5mC), a modification most commonly recognized as a stable, repressive regulator of gene expression. There is a large body of research that shows 5mC and other chemical modifications within DNA to have epigenetic roles in gene regulation. Identifying these marks and their function in biology is a fascinating area of epigenetics right now. ​

​​ 6. RNA modifications
Scientists are continually discovering new RNA modifications and new functions for existing modifications. Many RNA modifications thought only to exist in bacteria are being found in eukaryotic cells while others presumed only to exist on certain RNA, species such as tRNAs, are now being found to have crucial roles in mammalian mRNA translation. RNA modifications are very hot right now, and there is still a lot to explore in this field of research. ​​


Contents

H2AX becomes phosphorylated on serine 139, then called γH2AX, as a reaction on DNA double-strand breaks (DSB). The kinases of the PI3-family (Ataxia telangiectasia mutated, ATR and DNA-PKcs) are responsible for this phosphorylation, especially ATM. The modification can happen accidentally during replication fork collapse or in the response to ionizing radiation but also during controlled physiological processes such as V(D)J recombination. γH2AX is a sensitive target for looking at DSBs in cells. The presence of γH2AX by itself, however, is not the evidence of the DSBs. [5] The role of the phosphorylated form of the histone in DNA repair is under discussion but it is known that because of the modification the DNA becomes less condensed, potentially allowing space for the recruitment of proteins necessary during repair of DSBs. Mutagenesis experiments have shown that the modification is necessary for the proper formation of ionizing radiation induced foci in response to double strand breaks, but is not required for the recruitment of proteins to the site of DSBs.

DNA damage response Edit

The histone variant H2AX constitutes about 2-25% of the H2A histones in mammalian chromatin. [6] When a double-strand break occurs in DNA, a sequence of events occurs in which H2AX is altered.

Very early after a double-strand break, a specific protein that interacts with and affects the architecture of chromatin is phosphorylated and then released from the chromatin. This protein, heterochromatin protein 1 (HP1)-beta (CBX1), is bound to histone H3 methylated on lysine 9 (H3K9me). Half-maximum release of HP1-beta from damaged DNA occurs within one second. [7] A dynamic alteration in chromatin structure is triggered by HP1-beta release. This alteration in chromatin structure promotes H2AX phosphorylation by ATM, ATR and DNA-PK, [8] allowing formation of γH2AX (H2AX phosphorylated on serine 139). γH2AX can be detected as soon as 20 seconds after irradiation of cells (with DNA double-strand break formation), and half maximum accumulation of γH2AX occurs in one minute. [6] Chromatin with phosphorylated γH2AX extends to about a million base pairs on each side of a DNA double-strand break. [6]

MDC1 (mediator of DNA damage checkpoint protein 1) then binds to γH2AX and the γH2AX/MDC1 complex then orchestrates further interactions in double-strand break repair. [9] The ubiquitin ligases RNF8 and RNF168 bind to the γH2AX/MDC1 complex, ubiquitylating other chromatin components. This allows the recruitment of BRCA1 and 53BP1 to the long, modified γH2AX/MDC1 chromatin. [9] Other proteins that stably assemble on the extensive γH2AX-modified chromatin are the MRN complex (a protein complex consisting of Mre11, Rad50 and Nbs1), RAD51 and the ATM kinase. [10] [11] Further DNA repair components, such as RAD52 and RAD54, rapidly and reversibly interact with the core components stably associated with γH2AX-modified chromatin. [11] The constitutive level of γH2AX expression in live cells, untreated by exogenous agents, likely represents DNA damage by endogenous oxidants generated during cellular respiration. [12]

In chromatin remodeling Edit

The packaging of eukaryotic DNA into chromatin presents a barrier to all DNA-based processes that require recruitment of enzymes to their sites of action. To allow DNA repair, the chromatin must be remodeled.

γH2AX, the phosphorylated form of H2AX, is involved in the steps leading to chromatin decondensation after DNA double-strand breaks. γH2AX does not, itself, cause chromatin decondensation, but within 30 seconds of ionizing radiation, RNF8 protein can be detected in association with γH2AX. [13] RNF8 mediates extensive chromatin decondensation, through its subsequent interaction with CHD4, [14] a component of the nucleosome remodeling and deacetylase complex NuRD.

An assay for γH2AX generally reflects the presence of double-strand breaks in DNA, though the assay may indicate other minor phenomena as well. [15] On the one hand, overwhelming evidence supports a strong, quantitative correlation between γH2AX foci formation and DNA double-strand break induction following ionizing radiation exposure, based on absolute yields and distributions induced per unit dose. [15] On the other hand, not only the formation of distinct γH2AX foci but also the induction of pan-nuclear γH2AX signals have been reported as a cellular reaction to various stressors other than ionizing radiation. [16] The γH2AX signal is always stronger at DNA double-strand breaks than in undamaged chromatin. [16] γH2AX in undamaged chromatin is thought to possibly be generated via direct phosphorylation of H2AX by activated kinases, most likely diffusing from DNA damage sites.


A vision of 3D chromatin organization

DNA is wrapped around nucleosomes to form chromatin chains that can undergo further compaction to fit into the small space of a nucleus. The way chromatin is packaged in the 3D nucleus is crucial to its function however, owing to the technical challenge of visualizing chromatin in intact cells, what compact chromatin looks like in vivo has been the subject of debate for decades. O'Shea and colleagues have now developed a new imaging technique — ChromEMT (chromatin electron microscopy tomography) — that enables the visualization of both the local polymer structure and the global 3D organization of chromatin in the nucleus of intact interphase and mitotic human cells, challenging textbook models of chromatin organization.

Chromatin structure has been difficult to visualize in the nucleus with existing electron microscopy (EM) techniques owing to the lack of a high contrast and selective electron-dense stain that enables it to be distinguished from other components. The authors developed a DNA-labelling system (ChromEM), which uses a fluorescent DNA-binding dye (DRAQ5) that, upon excitation, catalyses the deposition of diaminobenzidine polymers on the surface of DNA, thus making it visible by EM. ChromEM in combination with multi-tilt EM tomography, in which large 3D cell volumes are imaged and reconstructed at multiple angles, enabled the direct visualization and reconstruction of chromatin structure and interactions inside the nucleus.

“chromatin in interphase nuclei was found to be organized into flexible and disordered chains”

The chromatin in interphase nuclei was found to be organized into flexible and disordered chains that range from 5 nm to 24 nm in diameter. Interestingly, this organization was seen throughout the nucleus, encompassing euchromatic and heterochromatic regions alike. Moreover, chromatin in mitotic chromosomes had a similar disordered structure and diameter range. This finding challenges the long-standing model of chromatin compaction whereby DNA-wrapped nucleosomes progressively fold into discrete higher-order chromatin fibres and ultimately into mitotic chromosomes. This hierarchical model predicts the formation of 30 nm (although there has been some controversy around this) and 120 nm fibres in interphase nuclei and of more compact 300 nm and 700 nm fibres in mitotic chromosomes.

Instead, O'Shea and colleagues propose that the overall primary structure of chromatin does not change. Different levels of compaction — that generate 3D nuclear domains in which DNA is more or less concentrated and thereby accessible — are achieved by bending flexible fibres at various lengths and creating contacts between and within chains. In mitotic chromosomes, the 3D concentration density of chromatin and such interactions is higher. This model is more plausible when considering the rapid dynamics of chromatin condensation that occur during mitosis.

In addition to prompting the revision of textbooks, ChromEMT opens up the possibility of studying how chromatin structure is linked to its function.


Watch the video: Chromosome chromatin and chromatid (October 2022).