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Can Euchromatin convert into Heterochromatin?

Can Euchromatin convert into Heterochromatin?


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I know that Heterochromatin can convert into Euchromatin but is the reverse possible? If yes, then How?


I would suggest in the future doing a small amount of research before asking. For example, the wikipedia page for euchromatin says this:

Euchromatin participates in the active transcription of DNA to mRNA products. The unfolded structure allows gene regulatory proteins and RNA polymerase complexes to bind to the DNA sequence, which can then initiate the transcription process. Not all euchromatin is necessarily transcribed, but in general that which is not is transformed into heterochromatin to protect the genes while they are not in use. There is therefore a direct link to how actively productive a cell is and the amount of euchromatin that can be found in its nucleus. It is thought that the cell uses transformation from euchromatin into heterochromatin as a method of controlling gene expression and replication, since such processes behave differently on densely compacted chromatin, known as the 'accessibility hypothesis'. One example of constitutive euchromatin that is 'always turned on' is housekeeping genes, which code for the proteins needed for basic functions of cell survival.

Facultative heterochromatin is chromatin which converts back and forth depending on circumstances. This occurs through (de)"condensation" or (de)"compaction" of chromatin:

Among the molecular components that appear to regulate the spreading of heterochromatin are the Polycomb-group proteins and non-coding genes such as Xist. The mechanism for such spreading is still a matter of controversy.[19] The polycomb repressive complexes PRC1 and PRC2 regulate chromatin compaction and gene expression and have a fundamental role in developmental processes.

I would suggest reading those pages a little bit and coming up with a more specific question if they don't answer it.


Difference Between Euchromatin and Heterochromatin

Our body is composed of billions of cells. A typical cell contains a nucleus, and the nucleus contains chromatin. According to biochemists, the operational definition of chromatin is the DNA, protein, RNA complex extracted from eukaryotic lysed interphase nuclei. According to them, the chromatin is the product formed from the packaged special proteins commonly known as histones. To put it simply, the chromatin is primarily the combination of deoxyribonucleic acid or simply DNA and other types of protein. Chromatin is the one responsible for packaging DNA into smaller volumes so that they can fit inside the cell. It is also responsible for strengthening the DNA for mitosis and meiosis to take place. Chromatin also prevents damaging the DNA and controls the gene expression and replication of the DNA.

There are two varieties of chromatin. They are euchromatin and heterochromatin. These two forms are distinguished in a cytological manner dealing with how intensely each form is stained. The euchromatin is less intense than heterochromatin. This only indicates that heterochromatin has tighter DNA packaging. To find out more about the difference between euchromatin and heterochromatin, this article will provide you a quick look regarding these two chromatin forms.

The lightly packed material is called euchromatin. Though it is lightly packed in the form of DNA, RNA, and protein, it is definitely rich in gene concentration and is usually under active transcription. If you are going to examine eukaryotes and prokaryotes, you will find the presence of euchromatin. Heterochromatin is only found in eukaryotes. When stained and observed under an optical microscope, euchromatin resembles light-colored bands while heterochromatin is dark colored. The standard structure of euchromatin is unfolded, elongated, and only about the size of a 10 nanometer microfibril. This minute chromatin functions in the transcription of DNA to mRNA products. The gene regulatory proteins, including the RNA polymerase complexes, are able to bind with the DNA sequence due to the unfolded structure of the euchromatin. When these substances are already bound, the transcription process begins. The activities of the euchromatin aid in cell survival.

On the other hand, heterochromatin is a tightly packed form of DNA. It is commonly found on the peripheral areas of the nucleus. According to some studies, there are probably two or more states of heterochromatin. Inactive satellite sequences are the main constituents of heterochromatin. The heterochromatin is responsible for gene regulation and protection of chromosomal integrity. These roles are made possible because of the dense DNA packing. When two daughter cells are divided from a single parent cell, heterochromatin is usually inherited, which means that the newly cloned heterochromatin contains the same DNA regions which results in epigenetic inheritance. There may be the occurrence of repression of transcribable materials due to the boundary domains. This occurrence may lead to the development of different levels of gene expression.

The following summary provides you a clearer understanding regarding the two forms of chromatin: euchromatin and heterochromatin.

Chromatin makes up the nucleus. It is made up of DNA and protein.

Chromatin has two forms: euchromatin and heterochromatin.

When stained and observed under an optical microscope, euchromatins are the light-colored bands while heterochromatins are the dark-colored bands.

Darker staining indicates tighter DNA packaging. Heterochromatins thus have tighter DNA packaging than euchromatins.

Heterochromatins are compactly coiled regions while euchromatins are loosely coiled regions.

Euchromatin contains less DNA while heterochromatin contains more DNA.

Euchromatin is early replicative while heterochromatin is late replicative.

Euchromatin is found in eukaryotes, cells with nuclei, and prokaryotes, cells without nuclei.

Heterochromatin is only found in eukaryotes.

The functions of euchromatin and heterochromatin are gene expression, gene repression, and DNA transcription.


Heterochromatin structure: lessons from the budding yeast

The eukaryotic genome can be roughly divided into euchromatin and heterochromatin domains that are structurally and functionally distinct. Heterochromatin is characterized by its high compactness and its inhibitory effect on DNA transactions such as gene expression. Formation of heterochromatin involves special histone modifications and the recruitment and spread of silencing complexes and causes changes in the primary and higher order structures of chromatin. The past two decades have seen dramatic advances in dissecting the molecular aspects of heterochromatin because of the identification of the histone code for heterochromatin as well as its writers and erasers (histone-modifying enzymes) and readers (silencing factors recognizing histone modifications). How heterochromatic histone modifications and silencing factors contribute to the special primary and higher order structures of heterochromatin has begun to be understood. The budding yeast Saccharomyces cerevisiae has long been used as a model organism for heterochromatin studies. Results from these studies have contributed significantly to the elucidation of the general principles governing the formation, maintenance, and function of heterochromatin. This review is focused on investigations into the structural aspects of heterochromatin in S. cerevisiae. Current understanding of other aspects of heterochromatin including how it promotes gene silencing and its epigenetic inheritance is briefly summarized.

Keywords: Saccharomyces cerevisiae Sir proteins heterochromatin higher order chromatin structure transcriptional silencing.


Key differences between Heterochromatin and Euchromatin

Following are the substantial points to differentiate among heterochromatin and euchromatin:

  1. The tightly packed form of DNA in the chromosome is called as heterochromatin, while the loosely packed form of DNA in the chromosome is called as euchromatin.
  2. In heterochromatin, the density of DNA is high and are stained dark, whereas in euchromatin the density of DNA is little and are lightlystained.
  3. Heterochromatin is found at the periphery of the nucleus in eukaryotic cells only, and Euchromatin is located in the inner body of the nucleus of prokaryotic as well as in eukaryotic cells.
  4. Heterochromatin shows little or no transcriptional activity as well they are genetically inactive, on the other hand, Euchromatin actively participates in the process of transcription and are genetically active also.
  5. Heterochromatin is compactly coiled and is late replicative, whereas Euchromatin is loosely coiled and early replicative.
  6. Regions of heterochromatin are sticky, but the areas of Euchromatin are non-sticky.
  7. In Heterochromatin part, the phenotype remains unchanged of an organism, though variation may be seen, due to the effect in DNA during the genetic process in the Euchromatin.
  8. Heterochromatin permits the gene expression regulation and also maintains the structural integrity of the cell though Euchromatin results in genetic variations, and allows the genetic transcription.

Conclusion

From the above information regarding chromatin – their structure and types. We can say that only Euchromatin is vigorously involved in the transcription process although heterochromatin and its types do not play such significant role.

Constitutive heterochromatin contains the satellite DNA, and it surrounds the centromere, and facultative heterochromatin is disbanded. So apparently it can be said that the eukaryotic cells and their inner structure are relatively complex.


Citation: Hughes SE, Hawley RS (2009) Heterochromatin: A Rapidly Evolving Species Barrier. PLoS Biol 7(10): e1000233. https://doi.org/10.1371/journal.pbio.1000233

Published: October 27, 2009

Copyright: © 2009 Hughes, Hawley. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.

Competing interests: The authors have declared that no competing interests exist.

Nearly 100 years ago, biologists divided regions of chromosomes into two types, euchromatin and heterochromatin, on the basis of their appearance (reviewed in [1]). The initial classification of DNA was based on the observation that euchromatic regions changed their degree of condensation during the cell division cycle, whereas heterochromatic regions remained highly condensed throughout the majority of the cell cycle. Although the biological significance of heterochromatin remained obscure for many years, it is now apparent that heterochromatin plays a number of biological roles, including a recently identified role in speciation. In addition to differences in the timing of chromosome condensation, numerous other differences have been identified between euchromatin and heterochromatin. Euchromatin is enriched with unique coding sequences, and the genes within the euchromatin are typically actively transcribed. Heterochromatin, on the other hand, is considered to be gene poor, being primarily composed of arrays of highly repetitive simple sequences, such as satellite sequences and/or transposable elements. Heterochromatin is enriched at the centromeres (see Figure 1) and telomeres of chromosomes.

Shown is a representative acrocentric chromosome containing both condensed heterochromatic (dark gray) and less condensed euchromatic regions (light gray). Beside each region are characteristics typical for each type of chromatin.

A number of chromatin modifications are associated specifically with either heterochromatin or euchromatin, such as specific methylation patterns on the histones. Proteins involved in creating the histone methylation patterns associated with heterochromatin, as well as proteins, such as heterochromatin protein 1 (HP1) that are involved in heterochromatin formation and gene silencing, are preferentially found localized to heterochromatic DNA. Finally, heterochromatin appears to be rapidly evolving, so that the sequence composition of heterochromatic regions from even closely related species is often distinct. The scarcity of genes in heterochromatin, as well as its rapid evolution, led many 20th century scientists to view heterochromatin as no more than “junk” DNA with little biological importance, other than comprising, and perhaps protecting, the centromeres and telomeres.

The fruit fly Drosophila melanogaster and its close relatives have been popular models for studying the nature and formation of heterochromatin (reviewed in [1]). In these species, euchromatin can easily be distinguished from pericentric heterochromatin, which surrounds the centromeres of mitotic chromosomes, by cytology owing to the differences in the condensation of the two types of DNA. Moreover, heterochromatic regions fail to replicate in the polytene chromosomes, which are highly replicated chromosomes that remain tightly associated in the larval salivary glands. This allows a reasonably precise demarcation of the junction between euchromatin and heterochromatin. Additionally, D. melanogaster is a highly tractable system for conducting genetic screens and for performing forward genetic manipulations. Because both heterochromatin and euchromatin are easily genetically dissected in D. melanogaster, the biology of heterochromatin has been intensely studied using this organism.

Indeed, work in D. melanogaster has begun to challenge the view of heterochromatin as “junk” DNA and demonstrated that heterochromatin plays a number of important cellular functions. The first indications of a “function” for heterochromatin came from studies of the multiple roles of heterochromatin in mediating recombination during meiosis [2],[3]. Perhaps more critically, placing normally euchromatic genes near heterochromatin causes the variable silencing of these genes, an effect known as position-effect variegation, or PEV (reviewed in [1]). This silencing effect and the fact that some genes must be in heterochromatin in order to be properly transcribed suggest that heterochromatin may play a critical role in the global control of gene regulation [4]–[6].

Moreover, experiments by Karpen, Dernburg, Hawley, and their collaborators have demonstrated that pairing of heterochromatic regions is required for the proper segregation of chromosomes that fail to undergo recombination during female meiosis [7]–[9]. Subsequent work demonstrated that chromosomes that fail to undergo recombination (the X and 4 th ) are connected by heterochromatic threads during prometaphase I in oocytes and that these threads are likely part of the mechanism by which heterochromatin facilitates nonrecombinant chromosome segregation [10].

Evidence for threads connecting chromosomes during meiosis in the sperm of both D. melanogaster and crane flies suggest a conserved function for thread-like structures in the segregation of chromosomes during meiosis [11],[12]. Although it has not yet been determined whether these threads are composed of heterochromatin, the repetitive intergenic spacer region of rDNA, which resides in heterochromatin, is required for the proper pairing of the Y chromosome with the X chromosome during male meiosis in D. melanogaster [13]. Finally, similar types of threads have been observed emanating from the heterochromatic centromere regions of chromosomes during mitosis in mammalian cells, suggesting that heterochromatic threads play an important role in chromosome segregation during mitosis as well [14],[15].

Given the critical functions of heterochromatic sequences in both meiosis and mitosis and its rapid change in sequence throughout evolution, it might not be surprising if differences in either heterochromatic sequences or the proteins that maintain them might indeed play a role in species isolation [16]. Mating of related species sometimes leads to the death or sterility of one or both sexes of progeny, which is known as hybrid incompatibility. Although hybrid incompatibility has been studied for decades, there have been only a few insights into the molecular mechanisms that underlie it, and in those cases known for the Drosophila species, the hybrid incompatibility has involved protein-coding genes [17],[18]. However, in this issue of PLoS Biology Ferree and Barbash demonstrate that the rapid divergence of heterochromatin also plays an important role in maintaining the reproductive isolation of D. melanogaster from the sister species Drosophila simulans [16]. The cross between D. simulans females and D. melanogaster males is unusual in that male offspring are viable but females die during embyronic development [19]. (Typically in cases of hybrid incompatibility the heterogametic males are the affected sex if only one sex is sterile or lethal.)

Ferree and Barbash found the lethality in hybrid female embryos resulted from failures during mitotic divisions 10–13 [16]. In these females, chromosomal regions frequently appeared highly stretched and lagged behind the other chromosomes during anaphase of these mitotic divisions [16]. The lagging DNA failed to become properly separated from its sister during mitosis, leading to improper chromosome segregation, aberrant mitotic divisions, and, ultimately, the death of female embryos.

Using fluorescent in situ hybridization, the authors determined that the lagging chromatin was primarily composed of the heterochromatic and highly repetitive 359-bp repeat on the X chromosome from the D. melanogaster father [16]. This particular heterochromatic repeat type has a different sequence composition and is much less abundant in D. simulans. The 359-bp repeat-containing region on the D. melanogaster X chromosome also overlaps the Zhr locus, a genetic region that was identified because its deletion from the D. melanogaster X chromosome allowed D. melanogaster males to produce viable hybrid daughters when crossed to D. simulans females [19].

The discovery that lethality in hybrid females results from a failure to maintain the integrity of a heterochromatic region of the D. melanogaster X chromosome containing the 359-bp repeat sequence suggests an intriguing possibility, namely, that the chromosome lagging and lethality of the D. melanogaster X chromosomal heterochromatin in female hybrids occurs because the D. simulans mother fails to provide a protein or RNA molecule required for proper maintenance and or separation during mitosis of the 359-bp repeat region provided by the D. melanogaster father (see Figure 2). One might even imagine that the failure in mitotic segregation observed here reflects a failure to properly resolve the heterochromatic threads observed to connect the pericentromeric heterochromatin in both mitotic and meiotic cells (see above).

This model is based on the results of the article published by Ferree and Barbash in this issue of PLoS Biology [16]. A cross between D. melanogaster males and D. simulans females, which results in hybrid females that die early in embryogenesis, is shown on the right. The drawing on the left depicts a cross between D. melanogaster males to D. melanogaster females for comparison. For simplicity, only the X chromosomes are shown and the heterochromatic region is specified with a darker color. In both crosses, the fusion of the sperm and egg results in zygotes carrying a pair of X chromosomes. The cross with the D. melanogaster females leads to normal chromosome segregation during anaphase of mitotic divisions 10–13 in female embryos. In the cross with D. simulans females mitosis fails to be completed normally in the hybrid female embryos. While the maternal X chromosomes segregate normally towards opposite spindle poles, the segregating centromeres of the paternally derived X chromosomes are connected by a bridge of chromatin. This bridge, which is heterochromatic and comprised of a region rich in the 359-bp repeat, causes improper segregation of the sister chromatids of the X chromosome, an event that eventually leads to aberrant mitotic divisions and ultimately the death of female hybrid embryos. The lagging or bridging of the 359-bp region is likely due to an absence of a maternally loaded factor in the D. simulans egg (shown as yellow diamonds in the D. melanogaster egg). We imagine that this factor might be involved in the resolution of heterochromatic threads that have been shown to connect the pericentromeric regions of both mitotic and meiotic chromosomes in normal cells (see text for a description). The absence of this factor prevents the proper formation or maintenance of chromatin structure in the 359-bp repeat region in female hybrid embryos.

The authors suspected that D. simulans lacks a factor required to either maintain heterochromatic stability or to resolve heterochromatic linkages in D. melanogaster X chromosomal heterochromatin during embryonic mitotic divisions. Indeed, they found that topoisomerase II, an enzyme required for proper mitosis, showed aberrant localization to the lagging DNA in hybrid embryos. Further work will be required to determine if other proteins or RNA molecules are absent or aberrantly localized from the D. simulans maternal cytoplasm, and if their absence is sufficient to cause the defects in the 359-bp repeat region in hybrid female embryos.

While this is the first example of a heterochromatic sequence causing hybrid incompatibility, other instances will likely be found in nature. Indeed, we cannot help but note a parallel between this example of hybrid inviability and a genetic phenomenon known as segregation distortion, which has also been well studied in D. melanogaster [20]. In this system a novel mutant known as SD, which is located in the euchromatin of Chromosome 2, prevents the meiotic transmission of homologous 2 nd chromosomes carrying high copy numbers of a heterochromatic element known as Responder (Rsp) [21]. The genetic basis of this phenomenon, which causes improper condensation and function of Rsp-bearing spermatids, is well understood, and a full molecular understanding of this process is within reach. While segregation distortion is indeed an example of “meiotic drive,” and not a species isolation mechanism, it bears mention here because it illustrates a second case in which a mutant in one strain impairs the function of a heterochromatic element in another thus illustrating the mechanisms of “heterochromatic incompatibility” may be more common than one might have expected.

As heterochromatin rapidly changes, the mechanisms that maintain it may well diverge as populations become isolated by various mechanisms. If those mechanisms change in such a way that the heterochromatin of population A can no longer be maintained by the maintenance proteins in population B, then the heterochromatin itself becomes a barrier between those populations as speciation proceeds. Many more questions await investigation, both in terms of the system of hybrid inviability described above and in terms of assessing the degree to which the safe-guarding of heterochromatic integrity underlies other examples of speciation. But one thing is clear: if any part of heterochromatin is indeed “junk,” then it is “junk” that both needs to be taken good care of and “junk” that sets one species apart from its neighbors.


Transcription organizes euchromatin similar to an active microemulsion

Chromatin is organized into heterochromatin, which is transcriptionally inactive, and euchromatin, which can switch between transcriptionally active and inactive states. This switch in euchromatin activity is accompanied by changes in its spatial distribution. How euchromatin rearrangements are established is unknown. Here we use super-resolution and live-cell microscopy to show that transcriptionally inactive euchromatin moves away from transcriptionally active euchromatin. This movement is driven by the formation of RNA-enriched microenvironments that exclude inactive euchromatin. Using theory, we show that the segregation into RNA-enriched microenvironments and euchromatin domains can be considered an active microemulsion. The tethering of transcripts to chromatin via RNA polymerase II forms effective amphiphiles that intersperse the two segregated phases. Taken together with previous experiments, our data suggest that chromatin is organized in the following way: heterochromatin segregates from euchromatin by phase separation, while transcription organizes euchromatin similar to an active microemulsion.


Abstract

Heterochromatin is a key architectural feature of eukaryotic chromosomes, which endows particular genomic domains with specific functional properties. The capacity of heterochromatin to restrain the activity of mobile elements, isolate DNA repair in repetitive regions and ensure accurate chromosome segregation is crucial for maintaining genomic stability. Nucleosomes at heterochromatin regions display histone post-translational modifications that contribute to developmental regulation by restricting lineage-specific gene expression. The mechanisms of heterochromatin establishment and of heterochromatin maintenance are separable and involve the ability of sequence-specific factors bound to nascent transcripts to recruit chromatin-modifying enzymes. Heterochromatin can spread along the chromatin from nucleation sites. The propensity of heterochromatin to promote its own spreading and inheritance is counteracted by inhibitory factors. Because of its importance for chromosome function, heterochromatin has key roles in the pathogenesis of various human diseases. In this Review, we discuss conserved principles of heterochromatin formation and function using selected examples from studies of a range of eukaryotes, from yeast to human, with an emphasis on insights obtained from unicellular model organisms.


Organisation of Chromosomes

Natella I. Enukashvily , Nikita V. Ponomartsev , in Advances in Protein Chemistry and Structural Biology , 2013

1 Introduction

In 1928, Heitz suggested the terms euchromatin and heterochromatin (HC) for differences detectable by suitable chromosomal stains ( Heitz, 1928 ). He stained cells from several species of moss with carmine acetic acid and observed a type of chromatin in the nucleus that remained condensed throughout the cell cycle. Heitz described that portion of the nuclear chromatin as heterochromatin, which maintained a condensed state (i.e., appeared darkly stained) throughout the cell interphase, while the remainder of the nuclear chromatin was extending to what he termed the euchromatin state. Cooper (1959) was able to summarize the data from Drosophila and suggested that heterochromatin and euchromatin differed in their biophysical conformations and in metabolic expression of their genes, but not in their basic structure of DNA arranged within chromosomes. Now, the concept of a eukaryotic genome consisting of two types of differently packed chromatin is widely accepted and it is included in school textbooks in biology. Although the term heterochromatin was originally defined cytologically as regions of mitotic chromosomes that remain condensed in the interphase, it is now more loosely applied to include regions of chromosomes that show characteristic properties such as, for example, gene repression and silencing (reviewed by Craig, 2005 Lohe & Hilliker, 1995 ).

It is commonly accepted that there are two types of heterochromatin—constitutive and facultative. The constitutive HC is thought to be condensed throughout the entire cell cycle unlike the facultative HC which is developmentally regulated. In mitotic chromosomes, constitutive heterochromatic regions are positioned most often in centromeric and pericentromeric regions as well as near telomeres. Chromosomes 1, 9, 16, and Y contain large blocks of heterochromatin. The facultative HC can be formed on different chromosomes regions. In mammalian females, one X-chromosome (either maternally or paternally derived) is randomly inactivated in early embryonic cells, with fixed inactivation in all descendant cells ( Lyon, 1961 ). Facultative HC can be formed in the promoter regions of non-transcribed genes ( Rand & Cedar, 2003 ). Some of the neocentromeres (regions of euchromatic DNA that acquired properties of centromeres) known to date can undergo heterochromatinization though they are formed within euchromatic regions ( Amor & Choo, 2002 Saffery et al., 2003 ).


Euchromatin Function

Despite being actively researched, the structure of chromatin is still poorly understood although it seems that the cycle in which the cell is at a certain time determines the structure of chromatin. Not surprisingly, the structure of euchromatin provides hints regarding its function and why it is present in transcriptionally active cells. As mentioned above, euchromatin is also called beads-on-a-string because of the resemblance between a necklace of beads connected through a string and the nucleosomes connected through the linker DNA. In this conformation, euchromatin is loose and consequently leaves the linker DNA exposed so that it can be transcribed this way, RNA and DNA polymerases as well as other proteins can access the DNA. Because of its loose structure, euchromatin is difficult to see under a microscope and appears faintly when stained—in contrast to the easily visible heterochromatin, which is densely packed.


MATERIALS AND METHODS

OI-DIC imaging system for biological samples

The principal schematic of the OI-DIC microscopy system is shown in Figure 1A. The microscope includes a light source with a bandpass filter, crossed linear polarizer and analyzer, phase shifter, condenser and objective lenses, tube lens, and charge-coupled device (CCD) digital camera. To overcome the limitations of conventional DIC imaging, the microscope contains two beam-shearing assemblies (Shribak, 2014). Each assembly consists of two identical DIC prisms and a 90° polarization rotator. Rotating light polarization by the rotators allows the shear directions to be rapidly switched by 90° without mechanically rotating the samples (e.g., live cells) or the prisms. We captured six raw images using OI-DIC microscopy, with two perpendicular shear directions and three biases ± 0.15λ and 0, where λ is the wavelength. The captured OI-DIC images were processed into OPD images (maps) whose intensity corresponded linearly to the OPD value. Image processing was performed using home-built software (OIDIC.exe). This and other processing algorithms were previously described (Shribak and Inoue, 2006 Shribak, 2013 Shribak et al., 2017).

To compare OI-DIC with the CellVista SLIM Pro phase-mapping microscope, manufactured by Phi Optics (Champaign, IL), we embedded 7-μm-diameter glass rods, which are used as spacers in liquid-crystal displays, in Fisher Permount mounting medium (Fisher Scientific, https://www.fishersci.com). The RIs of the glass rods and the Permount medium at wavelength 546 nm are 1.56 and 1.524, respectively. The images were obtained using 100 ×/1.4 NA oil immersion objective lenses. The CellVista SLIM captured and processed four phase-contrast images at the different biases.

Density estimation of cellular contents

To obtain calibration curves of the RIs of proteins and nucleic acids (Supplemental Figure S3), bovine serum albumin (BSA Sigma A-9418), and salmon sperm DNA (Fisher Scientific BP-2514) were dissolved in cell culture medium at concentrations of 0–200 and 0–30 mg/ml, respectively. The RIs of the prepared standard solutions were measured with a refractometer, Abbé-3L (Bausch & Lomb). The measured RIs and solution densities were plotted and fitted with linear functions to obtain calibration curves.

We estimated intracellular density distribution from obtained OPD maps using the following two steps (Figure 1B). First, we calculated the RI from the OPD. Because the OPD is proportional to the thickness of a sample and the difference in RI between the sample and the surrounding solution, as shown in Figure 1B, we calculated the RI of samples on the basis of the RI of the surrounding solution and sample thickness. Second, we obtained the dry mass density (“density” for short) of the sample from its RI, because the RI of a sample is proportional to its density. For proteins and nucleic acids, which are the dominant materials in mammalian cells (>60% of dry mass) (Alberts et al., 2007), our calibration curves (Supplemental Figure S3 see above) of RI versus dry mass density using BSA and salmon sperm DNA showed that both were well fitted to linear functions and were almost identical (RI = 1.3375 + 1.4 × 10 –4 × C, where C is dry mass density). Therefore, dry mass density in live cells, which consists mainly of proteins and nucleic acids, was calculated from their RI using a single calibration curve (Supplemental Figure S3). To estimate the densities of the total cell contents, we measured the average thickness of the cytoplasm and nucleus in each cell line (Supplemental Figure S1, A and B). The pericentric foci were assumed to be spherical (Supplemental Figure S1C). To obtain the RI of cytoplasm (RIcy), we used the RI of the surrounding culture medium (RImed, 1.3375) (Supplemental Figure S2). For the RIs of the nucleus and the pericentric foci, we used our calculated values of RIcy and RInuc, respectively (Supplemental Figure S2). These estimates were created using ImageJ software.

EGFP-MeCP2 construction

A plasmid containing MeCP2-EGFP was kindly provided by M.C. Cardoso (Brero et al., 2005). The moiety of MeCP2-EGFP was cut by XhoI and XbaI enzymes and blunted. The blunted fragment was inserted into the EcoRV site of the pEF5/FRT/VS-DEST Gateway Vector (Invitrogen) to generate pEF5-MeCP2-EGFP-FRT.

EGFP-mH3.1p-H3.1 construction

A mouse histone H3.1 promoter (∼840 base pairs)-H3.1 fragment was amplified from mouse embryonic stem (ES) cell genomic DNA by PCR using the following primer set:

5′-AGCCCTCTCCCCGCGGATGCGGCGGGCCAGCTGGATG­TCC-3′. The amplified fragment was cloned into an EGFP vector to obtain pmH3.1p-H3.1-EGFP. Then two more insert sequences were prepared. One was the H3.1promoter-H3.1-EGFP amplified from the pmH3.1p-H3.1-EGFP vector using the following primer set:

and 5′-TTACTTGTACAGCTCGTCCATGCCGAGAGT-3′. The other was the 3′ untranslated region of H3.1 (∼500 base pairs) amplified from the mouse ES genome using the primer pair: 5′-TGGACGAGC­TGTACAAGTAAAGTTCGTCTTTCTGTGTTTTTCAAA­GGCTC-3′

For backbone vector preparation, the region from the EF1α-promoter to the BGH polyadenylation signal sequence was removed from pEF5/V5-FRT Gateway (Invitrogen) to create the pFRT-Hygromycin vector. The pFRT-Hygromycin vector, H3.1promoter-H3.1-EGFP sequence, and 3′ untranslated region sequence were ligated by SLIC (Li and Elledge, 2007) to prepare the pmH3.1p-H3.1-EGFP-3′UTR-FRT plasmid. In addition, upstream of the H3.1 promoter, an insulator fragment (tandem cHS4 kindly provided by G. Felsenfeld) was inserted to obtain pIx2-mH3.1p-H3.1-EGFP-3′UTR-FRT.

EGFP-fibrillarin construction

To clone the fibrillarin gene, total RNA was isolated from NIH3T3 cells using the RNeasy Mini Kit (Qiagen), and first-strand cDNA was synthesized using the SuperScript III First-Strand Synthesis System (Thermo) with oligo(dT). The coding region of fibrillarin was amplified from the first-strand cDNA using the following primer pair: 5′-GGGGTACCATGAAGCCAGGTTTCAGCCC-3′ and 5′-GCGGGA­TCCTCAGTTCTTCACCTTGGGAG-3′. The amplified fragment was digested with KpnI and BamHI and ligated into KpnI/BamHI-digested pEGFP-C1 (Clontech, Palo Alto, CA). EGFP-fibrillarin fragments were excised, blunt-ended using T4 DNA polymerase (Takara, Japan), and inserted into EcoRV-precut pEF1-FRT to generate pEF1-EGFP-fibrillarin-FRT.

Cell culture and stable cell lines

We used RPE1, a human cell line, and NIH3T3, a mouse cell line. All of the cell lines were maintained in DMEM (Lonza) supplemented with 10% (vol/vol) fetal bovine serum (FBS) and 0.584 g/l l -glutamine (Sigma) at 37°C with 5% CO2 in air in a humidified incubator. To establish NIH3T3 cells expressing MeCP2-EGFP, EGFP-fibrillarin, or H3.1-EGFP, we used the Flp-In system (Invitrogen) as previously described (Maeshima et al., 2010 Hihara et al., 2012).

OI-DIC microscopy system

Details of the microcopy system are provided in the Supplemental Material.

Live-cell OI-DIC microscopy imaging

Cells were seeded on 24 mm × 24 mm square glass coverslips coated with poly d -lysine (Sigma) and cultured for 1–2 d. Then 30 min before imaging, 500 ng/ml Hoechst 33342 was added into the media and incubated further. The cells were mounted on a glass slide with a thin silicone spacer. We observed the mounted cells by OI-DIC and fluorescence imaging.

OI-DIC imaging of glass rods

A small number of glass rods 4 µm in diameter were suspended in two types of mineral oil with refractive indices of 1.54 and 1.58. Approximately 2 µl of the suspended solution was sandwiched between a glass slide and a coverslip, and then sealed with nail polish. The glass rods in the mineral oil were analyzed by OI-DIC microscopy using the same procedure as the live cell imaging.

Fixation

For formaldehyde fixation, cells on the square glass coverslips were washed once with phosphate-buffered saline (PBS), and then fixed in 4% formaldehyde at room temperature for 15 min. The fixed cells were washed with 10 mM HEPES-KOH, pH 7.5, 100 mM KCl, and 1 mM MgCl2 (HMK) (Maeshima et al., 2006) and permeabilized with 0.5% Triton X-100 in HMK at room temperature for 5 min. The treated cells were washed with HMK, stained with 500 ng/ml Hoechst 33342 in HMK at room temperature for 10 min, and then washed again with HMK. For MeOH fixation, cells on coverslips were fixed in ice-cold MeOH at –20°C for 30 min. Then the fixed cells were washed with HMK at room temperature for 15 min, stained with 500 ng/ml Hoechst 33342 in HMK for 10 min, and washed again with HMK. Finally, the cells were mounted on a glass slide and observed as described above.

Immunostaining of histone H3K9me3

Immunostaining was performed as previously described (Maeshima et al., 2010 Hihara et al., 2012). Cells were fixed in 2% formaldehyde (Wako) and permeabilized with Triton X-100. The primary and secondary antibodies were mouse anti-H3K9me3 (a generous gift from Hiroshi Kimura) and Alexa-Fluor-594-conjugated goat anti-mouse immunoglobulin G (Invitrogen) used at dilutions of 1:500 and 1:1000, respectively. Then DAPI (500 ng/ml) was added to the cells for 5 min, followed by washing with PBS prior to DNA staining. Images were obtained using a DeltaVision microscopy imaging system (Applied Precision) or a FLUOVIEW FV1000 confocal laser scanning microscope (OLYMPUS).

Measurements of cell thicknesses

Live RPE1 and NIH3T3 cells were seeded on 35 mm glass-bottom dishes. To fluorescently label DNA and cytoplasm, cells were incubated in culture medium containing 0.5 µg/ml Hoechst 33342 (Dojindo) and 5 µg/ml Calcein-AM (Dojindo) for 30 min. After washing out excess fluorescent dye, the stained cells were observed under an OLYMPUS FLUOVIEW FV1000 confocal laser scanning microscope equipped with a 60 ×/1.2 NA water objective. Hoechst 33342 and Calcein-AM fluorescence signals were acquired as three-dimensional image stacks (500 nm × 32 sections). The thicknesses of three regions (nucleus, cytoplasm, entire cell see Supplemental Figure S1) were measured in each cell line from the acquired stack images by ImageJ software.

Measurement of nuclear volume of live cells

Image stacks of live NIH3T3 and RPE1 cells acquired to measure cell thickness (described above) were used. The images were converted into binary images by auto thresholding in ImageJ software. Then the volumes of binary image stacks were analyzed using the 3D Objects Counter ImageJ plugin (Bolte and Cordelieres, 2006).

Quantification of fluorescence from Hoechst-stained DNA, MeCP2-EGFP, and α-H3K9me3

Image stacks of live or immunostained NIH3T3 cells (described above) were used. The middle sections of Hoechst-stained nuclei were selected from the z-stack images. The highest intensity of a focal plane of heterochromatin and the mean intensity of the surrounding low-intensity region in a nucleus were taken as the intensities of heterochromatin and the surrounding euchromatin. These values were adjusted to account for the background intensity.

Composition estimation of euchromatin and heterochromatin in live cells

The genome size of diploid mouse cells is 5.6 × 10 9 base pairs. Because 1 pg of DNA is 978 × 10 6 base pairs (978 × 10 6 base pairs/pg), the mass of the whole mouse genome is 5.73 pg. If we assume that nucleosomes (core histones + DNA) form every 200 base pairs of the genome and that the mass ratio of core histones to DNA is around 1:1, the mass of the nucleosomes in a mouse nucleus is 11.5 pg (5.73 × 2). The average mouse nuclear volume was calculated to be ∼1000 µm 3 (Supplemental Figure S6B), and the density of nucleosomes in mouse euchromatin was calculated to be 11.5 mg/ml. Further estimates are described under Results.

Monte Carlo simulation

Monte Carlo simulation is a computational algorithm that performs a numerical integration by making a random movement and evaluating whether the movement is acceptable based on the change in potential energy (Hibino et al., 2017). All of the molecules in the simulations were treated as hard spherical bodies. We employed a Metropolis Monte Carlo method without long-range potential or hydrodynamic interactions to determine the diffusive motion of molecules (Morelli and ten Wolde, 2008). The diameters and diffusion coefficients (Ds) of the crowding agents used in the simulations were 9.6 nm and 9 µm 2 s −1 , respectively, which are comparable to those of a single nucleosome molecule. The Ds of tracers (spheres with diameters of 5, 10, 15, and 20 nm) were 18, 9, 6, and 4.5 µm 2 s −1 , respectively. These Ds were determined by the Stokes–Einstein relationship based on parameters from the EGFP monomer, the diameter and D of which were 3.8 nm and 23.5 µm 2 s −1 , respectively (Hihara et al., 2012). Simulations were conducted in a 210-nm cubic box with two compartments (left and right halves) with periodic boundaries to avoid problems caused by finite space, and make the system more like an infinite one. For the “nucleosomes only” scenario, which corresponds to 11.5 mg/ml (euchromatin) and 85.9 mg/ml (heterochromatin), 134 and 968 copies of 9.6 nm spheres (crowding agents) were randomly placed in the left and right halves of the box, respectively. These crowding agents mimicked nucleosomes displaced less than 5 nm from their initial positions at t = 0 s (the “dog on a leash” model see also Hihara et al., 2012, and Maeshima et al., 2015). Then 50 tracers that could diffuse freely were placed in the left (euchromatin) region. The motion of the molecules was iteratively simulated following previously described procedures (Hihara et al., 2012 Maeshima et al., 2015). For the second simulation (nucleosomes + nonnucleosomal materials), 1578 and 1578–3945 9.6 nm spheres (crowding agents) were randomly placed in the left and right regions of the box, respectively, to represent euchromatin and heterochromatin (1.53–2.5-fold density differences). To represent nucleosomes, 134- and 968 9.6-nm spheres were randomly placed in the left and right regions, with their behavior following the “dog on a leash” model. The rest of the spheres moved freely only in each half, to represent diffusing proteins and RNAs. Then 50 tracers were placed in the left (euchromatin) region. Although the simulation process was similar to the first simulation, we restricted the movements of crowding agents to within their regions to keep the density of each region constant. Results were obtained by averaging 150 samples from three independent trials. The simulation time step, Δt, was 10 ns.

Statistical analyses

All of the statistical analyses were performed using the two-tailed Student’s t test. p values less than 0.05 were considered statistically significant.


Watch the video: DNA Packaging Animation. chromatin, histone and nucleosome modifications (July 2022).


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