Why are Chromosome Territories important?

Why are Chromosome Territories important?

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Chromosomes occupy discrete regions of the nucleus, referred to as 'Chromosome Territories'. This spatial organization is emerging as a crucial aspect of gene regulation and genome stability in health and disease.

But how is this the case when:

Most eukaryotes seem to have Chromosome Territories, but yeast S. cerevisiae is an exception to this.

So if some eukaryotes do, and some do not, what information do Chromosome Territories provide? Some literature uses the phrase 'non-random chromosome territory arrangements'. Thus, I am confused.

Chromosome territories are randomly arranged in some eukaryotes.

How are chromosome territories crucial for gene regulation and genome stability in health and disease in some eukaryotes, but random in other eukaryotes?

First, this idea was discredited or ignored during several decades, some studies suggest that CT is random in some eukaryotes that isn't necessary the truth. It is not easy at all to determine is CT are random or not

Second, not all eukaryotic organism are the same. Plants and Fungi have cell walls, animals do not. Fungi can have a dikaryotic cell or even undergo somatic hybridisation.

Third, "chromosomes are more unfolded in lower eukaryotes such as yeast."Chromosome Territories: The Arrangement of Chromosomes in the Nucleus

This article does not mention that S. cerevisiae is an exception, on the contrary "and may even hold for single-cell eukaryotes, such as budding and fission yeast".

Thank you for the question, I learnt a lot while searching for an answer


A chromosome is a long DNA molecule with part or all of the genetic material of an organism. Most eukaryotic chromosomes include packaging proteins called histones which, aided by chaperone proteins, bind to and condense the DNA molecule to maintain its integrity. [1] [2] These chromosomes display a complex three-dimensional structure, which plays a significant role in transcriptional regulation. [3]

Chromosomes are normally visible under a light microscope only during the metaphase of cell division (where all chromosomes are aligned in the center of the cell in their condensed form). [4] Before this happens, each chromosome is duplicated (S phase), and both copies are joined by a centromere, resulting either in an X-shaped structure (pictured above), if the centromere is located equatorially, or a two-arm structure, if the centromere is located distally. The joined copies are now called sister chromatids. During metaphase the X-shaped structure is called a metaphase chromosome, which is highly condensed and thus easiest to distinguish and study. [5] In animal cells, chromosomes reach their highest compaction level in anaphase during chromosome segregation. [6]

Chromosomal recombination during meiosis and subsequent sexual reproduction play a significant role in genetic diversity. If these structures are manipulated incorrectly, through processes known as chromosomal instability and translocation, the cell may undergo mitotic catastrophe. Usually, this will make the cell initiate apoptosis leading to its own death, but sometimes mutations in the cell hamper this process and thus cause progression of cancer.

Some use the term chromosome in a wider sense, to refer to the individualized portions of chromatin in cells, either visible or not under light microscopy. Others use the concept in a narrower sense, to refer to the individualized portions of chromatin during cell division, visible under light microscopy due to high condensation.

Chromosome Break☆

Wendy J. Cannan , David S. Pederson , in Reference Module in Life Sciences , 2017

4 How are chromosome breaks repaired?

Unwanted chromosome breaks are considered the single most deleterious form of DNA damage to the cell. The two main pathways to repair of double-strand breaks, non-homologous end-joining (NHEJ) and DNA homology-mediated recombination repair (HRR or HR), are described elsewhere in much greater detail. Here, we note only that (outside of meiosis) HR uses the genetic information from the sister chromatids as templates for repair, and is relatively error-free. Hence, DNA breaks that occur during G1 or early S phase cannot be repaired via HR. Such breaks are instead channeled into the NHEJ pathway, which directly joins two broken DNA ends. These joining events entail DNA ligation reaction that can occur only if the DNA substrate contains a 5′-phosphate group, and a 3′-hydroxyl group. “Clean” DNA breaks, such as those produced by restriction endonucleases, can be efficiently ligated without the participation of other factors. However, DNA breaks produced during reactions between ROS and DNA are rarely clean. DNA moieties at or near IR-generated breaks may include 3′-phosphates, 3′-phosphoglycolates, 5′-aldehydes, and 5′-hydroxyls. These may slow or inhibit double-strand break repair ( Weinfeld and Soderlind, 1991 ). No single enzyme can remove all of these moieties, and DNA end processing prior to ligation may sometimes take hours (reviewed in Andres et al. (2015) ). As well, the DNA processing needed to produce ligatable ends often produce short deletions, making NHEJ mutagenic.


Figure (PageIndex<3>): Overview of Meiosis. During meiosis, homologous chromosomes separate and go to different daughter cells. This diagram shows just the nuclei of the cells. Notice the exchange of genetic material that occurs prior to the first cell division.

The process that produces haploid gametes is called meiosis. Meiosis is a type of cell division in which the number of chromosomes is reduced by half. It occurs only in certain special cells of an organism. In mammals, Meiosis occurs only in gamete producing cells within the gonads. During meiosis, homologous (paired) chromosomes separate, and haploid cells form that have only one chromosome from each pair. Figure (PageIndex<3>) gives an overview of meiosis.

As you can see from the meiosis diagram, two cell divisions occur during the overall process, so a total of four haploid cells are produced. The two cell divisions are called meiosis I and meiosis II. Meiosis I begins after DNA replicates during interphase. Meiosis II follows meiosis I without DNA replicating again. Both meiosis I and meiosis II occur in four phases, called prophase, metaphase, anaphase, and telophase. You may recognize these four phases from mitosis, the division of the nucleus that takes place during routine cell division of eukaryotic cells.

Figure (PageIndex<4>): Complete Stages of Meiosis: An animal cell with a diploid number of four (2n = 4) proceeds through the stages of meiosis to form four haploid daughter cells.

Meiosis I

  1. Prophase I: The nuclear envelope begins to break down, and the chromosomes condense. Centrioles start moving to opposite poles of the cell, and a spindle begins to form. Importantly, homologous chromosomes pair up, which is unique to prophase I. In prophase of mitosis and meiosis II, homologous chromosomes do not form pairs in this way. During prophase I, crossing-over occurs. The significance of crossing-over is discussed in the next section called variations.
  2. Metaphase I: Spindle fibers attach to the paired homologous chromosomes. The paired chromosomes line up along the equator of the cell. This occurs only in metaphase I. In metaphase of mitosis and meiosis II, it is sister chromatids that line up along the equator of the cell.
  3. Anaphase I: Spindle fibers shorten, and the chromosomes of each homologous pair start to separate from each other. One chromosome of each pair moves toward one pole of the cell, and the other chromosome moves toward the opposite pole.
  4. Telophase I and Cytokinesis: The spindle breaks down, and new nuclear membranes form. The cytoplasm of the cell divides, and two haploid daughter cells result. The daughter cells each have a random assortment of chromosomes, with one from each homologous pair. Both daughter cells go on to meiosis II.

Meiosis II

  1. Prophase II: The nuclear envelope breaks down and the spindle begins to form in each haploid daughter cell from meiosis I. The centrioles also start to separate.
  2. Metaphase II: Spindle fibers line up the sister chromatids of each chromosome along the equator of the cell.
  3. Anaphase II: Sister chromatids separate and move to opposite poles.
  4. Telophase II and Cytokinesis: The spindle breaks down, and new nuclear membranes form. The cytoplasm of each cell divides, and four haploid cells result. Each cell has a unique combination of chromosomes.

Yeast and Cancer

Over the last several decades, researchers have been tirelessly interrogating all of the mutations that cause cancer in humans. Dr. Leeland Hartwell, a biologist and 2001 Nobel Laureate, was one of the first scientists to discover some of the mutations involved in cancer. Since then, many of the mutations found so far are in genes involved in some way with cell division and DNA replication. And in many of these cases, as with Dr. Hartwell, these mutations have been found in other organisms, like yeast, before their relevance in human cancer was realized. Through Dr. Hartwell’s work, he found that genes involved in the “cell division cycle” (CDC) in S. cerevisiae, were also found in a similar capacity in humans. Over his career, he identified over 100 genes involved in the control of cell division. The discoveries made by Dr. Hartwell and others using yeast as a model organism have contributed significantly to our understanding of how cell division is controlled in eurkaryotic cells. This understanding has had broad applications across many biological fields including the prevention, diagnosis, and treatment of cancer.

Learn more about the impact of Dr. Hartwell's discoveries in this article published by Fred Hutch.

Materials and Methods

Cells, fixation procedure, and 3D FISH pretreatment.

A vigorously growing primary human fibroblast culture was established from a skin biopsy of a 2-y-old boy. Chromosome banding and M-FISH analyses performed after the second passage (1:2 split) showed a normal male karyotype (46, XY). Surplus cultures were kindly provided by the Abteilung Medizinische Genetik, Munich University, Germany cells were further grown in our laboratory in DMEM medium supplemented with 10% FCS, and aliquots were frozen at about 5–7 passages in liquid nitrogen. Cells from these aliquots were further propagated for 2–4 wk, subcultivated (1:2) every 5 d, and routinely checked for absence of mycoplasm contaminations [56]. For 3D FISH and ReFISH experiments, cells were seeded on coverslips (26 × 76 mm, thickness 0.17 ± 0.01 mm). For studies of quiescent cell populations, cultures were grown to confluence and maintained for 1 wk before 3D fixation was performed in 4% paraformaldehyde/1× PBS for 10 min [30]. Control experiments with BrdU pulse labeling (1 h) and immunostaining of the cell-cycle-specific nuclear protein Ki67 indicated that more than 99.5% of the cells were in a quiescent state (G0) under these conditions. To investigate G0 and S-phase cells simultaneously on the same coverslip, cells were fixed at approximately 40%–50% confluence. Nuclei from cells in and out the cell cycle were discriminated by BrdU pulse labeling (45 min) and pKi67 staining. Growing fibroblast cultures were also used to obtain prometaphase rosettes. Nuclei in S-phase showed pKi67 staining and incorporation of BrdU, while G0 nuclei lacked both signals (data not shown). An amniotic fluid cell culture from a female fetus (46, XX) was established following diagnostic amniocentesis, and a growing early-passage culture was pulse-labeled with BrdU (45 min) prior to 3D fixation. Permeabilization steps performed prior to 3D FISH included treatments with 0.5% Triton-X100 (20 min), 20% glycerol in PBS (30 min), repeated freeze/thawing in liquid nitrogen, and incubation in 0.1 M HCl (5 min) or pepsin (0.002% in 0.01 M HCl). Slides were stored at 4 °C in 50% formamide/2× SSC until 3D FISH was performed.

DNA probes, labeling protocols, 3D FISH, and probe detection.

Whole chromosome painting probes were kindly provided by Malcolm Ferguson-Smith (Cambridge University, United Kingdom). Probes were established from flow-sorted human chromosomes and amplified by DOP-PCR.

For 24-color 3D FISH experiments in a single assay, chromosome paint probes for all 24 chromosome types (HSAs 1–22, X, and Y) were labeled using a combinatorial labeling scheme with seven differentially labeled nucleotides including diethylaminocoumarine (DEAC NEN Life Science Products, Zaventem, Belgium), Fluorogreen (Amersham Pharmacia Biotech, Piscataway, New Jersey, United States), TexasRed (Molecular Probes, Eugene, Oregon, United States), Cy3, Cy5 (Amersham Pharmacia Biotech), biotin-dUTP (Rockland Immunochemicals, Gilbertsville, Pennsylvania, United States) and digoxigenin-dUTP (Roche, Basel, Switzerland) [31]. A mix containing the 24 labeled probes in 50% formamide/10% dextran sulfate/1× SSC was hybridized for 3 d at 37 °C as previously described [30]. Post-hybridization washes were performed three times with 0.5× SSC at 60 °C. Avidin-Cy5.5 was used for detection of biotin and anti-dig-Cy7 (custom-made) for the immunodetection of digoxigenin.

For the differential coloring of all 24 chromosome types in ReFISH experiments [30,32], chromosome-specific paint probes from all 24 human chromosome types were labeled with either biotin-dUTP, TAMRA-dUTP, or Spectrum Green–dUTP. Two different hybridization mixtures were prepared in a way that allowed the unequivocal discrimination of each chromosome type in two subsequent hybridization experiments. After hybridization of the first probe subset, biotinylated probes were detected by Avidin-Cy5.5. Confocal image stacks were acquired as described below, and the cell coordinates were recorded. Thereafter, the same cells were re-hybridized with the second probe subset, followed again by detection with Avidin-Cy5.5 and confocal microscopy.

In two-color 3D FISH experiments aimed at the simultaneous visualization of two pairs of homologous CTs, we used biotin- and digoxigenin-labeled chromosome paint probes for HSAs 18/19, HSAs 17/Y, and HSAs 1/20.


After 3D FISH of all 24 chromosome types in a single assay, nuclei were imaged with an epifluorescence wide-field microscope (DMRXA, Leica, Wetzlar, Germany), equipped with a Plan Apo 63×/1.4 oil immersion objective, an 8-filter wheel with narrow-band filters (Chroma Group, San Bruno, California, United States), and an automated z-step motor [57]. For image capturing, a Sensys CCD camera (PhotoMetrics, Huntington Beach, California, United States) was used. Both camera and microscope were controlled by Leica QFluoro software. Stacks of optical sections with an axial distance of 250 nm were collected from nuclei with a regular shape showing apparently complete and specific hybridization signals in all channels. For each optical section images were collected sequentially for all fluorochromes. Stacks of 8-bit gray-scale 2D images were obtained with a pixel size of 110 nm in the x and y directions, and an image size of 256 × 256 pixels.

In ReFISH experiments, images for the three fluorochromes were obtained from the same nuclei after the first and second hybridization with a Zeiss (Oberkochen, Germany) LSM 410 confocal microscope equipped with a 63× Plan Apo objective and filters for FITC, Cy3, and Cy5. Scans were sequentially performed for the three fluorochromes on each light-optical section. An alignment of the two image stacks obtained for each nucleus after the first and second hybridization was performed on a Silicon Graphics (Mountain View, California, United States) workstation (OS Irix 6.2) using the program Correlator [58] in the integrated development environment Khoros (Khoral, Albuquerque, New Mexico, United States). This procedure allowed the fitting of the two image stacks with subvoxel accuracy. A comparison of the nuclear shape after the first and second hybridization did not reveal a notable difference, but we observed a slight increase in volume, which was corrected by the computer algorithm.

Deconvolution and image processing.

Each fluorochrome channel was normalized to a maximum intensity value of 255, and subjected to deconvolution by the software Huygens (Scientific Volume Imaging, Hilversum, The Netherlands). Thereafter, chromosomes in the image stacks were classified according to their labeling scheme using the software goldFISH [33] running on a Silicon Graphics workstation. This software carried out an automated classification of fluorescently stained areas in each light-optical nuclear section on the basis of the combinatorial labeling scheme. A false color representing the classified chromosome type was allocated to each classified territory. The software calculated the 3D IGCs of each classified territory as well as the CN or the CR by means of the DAPI image stack. Maximum intensity projections of image stacks were made with ImageJ software (National Institutes of Health, Bethesda, Maryland, United States). Displayed overlays were processed with Adobe Photoshop (Adobe Systems, San Jose, California, United States). Three-dimensional reconstructions of image stacks were performed using Amira 2.3 (Mercury Computer Systems, Chelmsford, Massachusetts, United States). Overlap of CTs in nuclei or of chromosomes in 3D fixed mitotic rosettes can be a source of misclassification. Because of the remarkable flatness of fibroblast nuclei in G0 (maximum height approximately 6 μm, with CTs often expanded from the bottom of the nucleus to the top), errors due to CT overlaps are less likely than in spherical nuclei.

Data evaluation.

For each classified PC and CT we determined its 3D IGC, together with the CR and the CN, respectively. IGC coordinates were imported into Excel (Microsoft, Redmond, Washington, United States), and the following distances and angles were measured: (1) 3D CR–PC and 3D CN–CT distances (3D radial distances), (2) 3D PC–PC and 3D CT–CT distances between all possible pairs of homologous and heterologous PCs or CTs, and (3) 3D PC–CR–PC and 3D CT–CN–CT angles. For comparison of different nuclei, 3D distances were normalized using the following procedure. A coordinate system was applied to each individual nucleus with the nuclear center as the origin (polar coordinates). The angle α between the longer cell axis and the x-axis of the coordinate system was determined, and the coordinates were recalculated following a rotation: x′ = cosα x − sinα y and y′ = sinα x − cosα y. For size measurement of nuclei, the DNA was stained with DAPI or TOPRO-3. The diameters in X and Y were then measured using the light-optical section with maximum lateral nuclear expansion. The height was measured between the upper and lower plane showing the most peripheral DAPI staining along the z-axis. The relative radial distance r of a PC or CT was calculated as r = (r1/r0)·100, where r1 represents the distance CN–CT or CR–PC, respectively, and r0 denotes either the distance between the CR and the prometaphase edge or between the CN and the nuclear edge, drawing a line through the IGC of the analyzed chromosome. Angles were calculated between the IGCs of homologous CTs or PCs using the CN or CR as the midpoint. The relative radial distance between heterologous PCs or CTs was calculated as a fraction of the nuclear diameter.

Distance and angle measurements obtained for IGCs of CTs and PCs were compared with distances and angles calculated between points statistically placed by a random number generator (“random point distribution model”).

MDS plots were generated by SPSS 11 (SPSS, Chicago, Illinois, United States). This program provided a 2D distance map taking into account the normalized mean heterologous 3D CT–CT distances calculated for all possible combinations of heterologous CTs. All distances were normalized to the diameter of the nucleus before generating the plots. The units after the transformation to a MDS map are arbitrary since the distances are merely relative. For a quantitative 3D evaluation of CT distributions in two-color 3D FISH experiments, the 3D-RRD computer program was used (see [25] and [59] for detailed description). Briefly, the program determines (1) the center of gravity of a given nucleus and its borders, on the basis of the DNA counterstain, and (2) all voxels of painted chromosome territories. The nuclear radius in any direction from the nuclear center of gravity to the segmented nuclear edge was normalized to 100%, and the nuclear space was divided into 25 concentric shells. All shells possessed the same thickness along each possible radial vector from the center of the nucleus to the periphery. Accordingly, the thickness of these shells in flat ellipsoidal nuclei was much larger along the x- and y-axis than along the z-axis. Thus, the distribution of the DNA of painted CTs was measured and expressed as a function of the relative distances of each shell from the center of the nucleus. Significance tests were carried out with either SPSS 11 or Sigma Stat (SPSS). If not otherwise stated a K-S significance test was applied.

Modeling of human fibroblast cell nuclei with statistical CT distributions.

To simulate the statistical distribution of CTs in human fibroblast nuclei, the SCD model was applied [25,41,60]. The DNA content of individual human chromosomes [61] was used to estimate the number of 1-Mbp chromatin domains constituting a given model chromosome. These domains were represented by 500-nm diameter spheres. Model nuclei were generated with an ellipsoidal shape with half-axes representing the average half-axes of human G0 fibroblast nuclei determined from light-optical stacks (x = 10 μm, y = 5 μm, and z = 2.5 μm).

Briefly, starting configurations representing a statistical chromatid distribution in male diploid human fibroblasts (46, XY) at late anaphase/telophase were established as follows. The location of the center of gravity of each chromatid was initially represented by the mass center of a small inelastic sphere (iS). The volume of a given iS was proportional to the DNA content of its natural chromosome counterpart, while its radius was very small compared with the half-axes of the ellipsoid. The total volume of the 46 iSs (representing the 46 chromosomes of the diploid human complement) comprised 22% of the total volume of the ellipsoidal model nucleus. The mass centers of the 46 iSs were statistically placed into the ellipsoid as follows. Using a random-number generator, the 3D coordinates for the mass centers of all iSs were generated iteratively in a nonoverlapping fashion. If the addition of a new iS yielded any overlap with the position of already existing iS, the 3D coordinates were discarded and new randomly generated 3D coordinates were tested. This process was repeated until all 46 iSs were located in the model nucleus. In a second step, chromatids were modeled as small rods with a spherical cross section of 500 nm (see Figure 1D). Each rod represents a linear chain of spherical 1-Mbp chromatin domains. To model the dense packaging of 1-Mbp domains in chromatids, a distance of 13 nm was simulated between the centers of gravity of any two adjacent 1-Mbp domains along the chromatid axis. For example, a rod representing a chromatid with a DNA content of 100 Mbp had a length of 1,300 nm. In this way we modeled highly compacted and rigid chromatids. Rods were placed with a random orientation inside the 46 iSs in such a way that their centers of gravity coincided with the centers of the iSs from step one. As a third step, the relaxation of the statistically placed chromatids into decondensed CTs was simulated. For this purpose Monte Carlo relaxation loops were carried out with about 400,000 steps to obtain thermodynamic equilibrium configurations [14]. For decondensed CTs we assumed 120-kb linker connections between neighboring 1-Mbp chromatin domains, representing higher-order clusters of 100-kb loop domains [62]. These connections were modeled by entropic spring potentials enforcing a mean distance of 600 nm between the centers of gravity of adjacent domains. For distances between the centers of two adjacent 1-Mbp domains of 500 nm or more, we assumed that their repulsive potential was zero. When distances became smaller than 500 nm, the repulsive potential became increasingly positive, resulting in an increasing mutual repulsion between the two domains. Long-term Monte Carlo relaxation loops showed that the two assumptions of a spring potential and a repulsive potential are not sufficient to maintain the experimentally observed compactness of CTs. To achieve model CTs with diameters comparable to CTs in fibroblast nuclei, we introduced a weak potential barrier around each simulated chromatin domain chain representing a given chromosome. In each Monte Carlo step the 3D coordinates of a randomly chosen 1-Mbp domain were changed slightly for each CT. The new coordinates were accepted if the resulting chromatin domain configuration came closer to the thermodynamic equilibrium. Otherwise they were rejected and the process repeated. The 3D positions of the CT centers of gravity after 400,000 Monte Carlo steps represented the statistical arrangement of CTs in model nuclei. The further evaluation of distances and angles between model CTs was performed as described for experimental fibroblast nuclei.

Polymer models and cell-to-cell variability in chromatin folding

Different polymer models can also be distinguished by their prediction of cell-to-cell variability of chromatin folding. A polymer model describes the ensemble of the folding configurations of the polymer and each of these conformations describes one possible geometric structure of the polymer. The configuration of the polymer statistically varies over time. This translates to a cell-to-cell variability that can be measured in a population of fixed cells – because fixation creates a snapshot of every cell – each in a different chromatin folding configuration at the time of fixation. For example, the RW polymer model predicts that the physical distance R between two defined monomers of the polymer, when measured in many cells, shows a Gaussian distribution. In general, distributions are characterised by their moments, a set of parameters that uniquely characterises the distribution. For example, the first moment is the mean value of the distribution, the second moment is its width (variance). The higher moments describe other features of the distribution. Here, the ratio between the fourth moment of the distribution and the second moment squared is of interest, as it gives a dimensionless number that is independent of any parameter that can be fitted to the experimental data. Using these moments, experimentally obtained distributions – when measured with sufficiently accuracy – can be compared with polymer model predictions. For instance, a set of 3D FISH measurements obtained from a large number of individual fixed cells reflects the cell-to-cell variation and yields a distribution of the physical distance R as a function of the genomic distance (Box 1). For this distribution one can compute the moments and compare the experimental data to model predictions. Such a comparison has shown that linear polymer models, i.e. models not involving loops, are incompatible with the experimental data (Bohn and Heermann, 2009).

Relationship between physical distance and genomic distance for linear and looped polymer models. (A) Schematic representation of the physical distance R between two points on an interphase chromosome and the genomic distance g. (B) Linear polymer model that does not assume looping. This model predicts that the mean squared distance ⟨R 2 ⟩ increases linearly with increasing genomic distance g. (C) Model that assumes a looped chromatin chain. In contrast to the linear model, looped models predict that ⟨R 2 ⟩ reaches a plateau at large genomic distances – as it has been found experimentally (Mateos-Langerak et al., 2009).

Relationship between physical distance and genomic distance for linear and looped polymer models. (A) Schematic representation of the physical distance R between two points on an interphase chromosome and the genomic distance g. (B) Linear polymer model that does not assume looping. This model predicts that the mean squared distance ⟨R 2 ⟩ increases linearly with increasing genomic distance g. (C) Model that assumes a looped chromatin chain. In contrast to the linear model, looped models predict that ⟨R 2 ⟩ reaches a plateau at large genomic distances – as it has been found experimentally (Mateos-Langerak et al., 2009).

Chromosomes and Cell Division

One of the most important elements of successful cell division is the correct distribution of chromosomes. In mitosis, this means that chromosomes must be distributed between two daughter cells. In meiosis, chromosomes must be distributed among four daughter cells. The cell's spindle apparatus is responsible for moving chromosomes during cell division. This type of cell movement is due to interactions between spindle microtubules and motor proteins, which work together to manipulate and separate chromosomes.

It is vitally important that a correct number of chromosomes be preserved in dividing cells. Errors that occur during cell division may result in individuals with unbalanced chromosome numbers. Their cells may have either too many or not enough chromosomes. This type of occurrence is known as aneuploidy and may happen in autosomal chromosomes during mitosis or in sex chromosomes during meiosis. Anomalies in chromosome numbers can result in birth defects, developmental disabilities, and death.

The demise of men?

As we argue in a chapter in a new e-book, even if the Y chromosome in humans does disappear, it does not necessarily mean that males themselves are on their way out. Even in the species that have actually lost their Y chromosomes completely, males and females are both still necessary for reproduction.

In these cases, the SRY “master switch” gene that determines genetic maleness has moved to a different chromosome, meaning that these species produce males without needing a Y chromosome. However, the new sex-determining chromosome – the one that SRY moves on to – should then start the process of degeneration all over again due to the same lack of recombination that doomed their previous Y chromosome.

However, the interesting thing about humans is that while the Y chromosome is needed for normal human reproduction, many of the genes it carries are not necessary if you use assisted reproduction techniques. This means that genetic engineering may soon be able to replace the gene function of the Y chromosome, allowing same-sex female couples or infertile men to conceive. However, even if it became possible for everybody to conceive in this way, it seems highly unlikely that fertile humans would just stop reproducing naturally.

Although this is an interesting and hotly debated area of genetic research, there is little need to worry. We don’t even know whether the Y chromosome will disappear at all. And, as we’ve shown, even if it does, we will most likely continue to need men so that normal reproduction can continue.

Indeed, the prospect of a “farm animal” type system where a few “lucky” males are selected to father the majority of our children is certainly not on the horizon. In any event, there will be far more pressing concerns over the next 4.6m years.

Is the genome now completely sequenced?

Well, no. An obvious omission is the Y chromosome, because the complete hydatidiform mole cells used to compile this sequence contained two identical copies of the X chromosome. However, this work is underway and the researchers anticipate their method can also accurately sequence the Y chromosome, despite it having highly repetitive sequences.

Even though sequencing the (almost) complete genome of a human cell is an extremely impressive landmark, it is just one of several crucial steps towards fully understanding humans’ genetic diversity.

The next job will be to study the genomes of diverse populations (the complete hydatidiform mole cells were European). Once the new technology has matured sufficiently to be used routinely to sequence many different human genomes, from different populations, it will be better positioned to make a more significant impact on our understanding of human history, biology and health.

Both care and technological development are needed to ensure this research is conducted with a full understanding of the diversity of the human genome to prevent exacerbation of health disparities by limiting discoveries to specific populations.

Watch the video: BY437 0129 14 - Chromosome territories (August 2022).