Information

9: Changes in Chromosome Number and Structure - Biology


Previous chapters described chromosomes as simple linear DNA molecules on which genes are located. For example, your largest chromosome, chromosome 1, has about 3536 genes. To ensure that each of your cells possesses these genes the chromosome has features that allow it to be passed on during cell division. Origins of replication found along its length provide places for DNA replication to start, telomeres protect each end of the chromosome, and a single centromere near the middle provides a place for microtubules to attach and move the chromosome during mitosis and meiosis. This chapter examines: (1) changes in the number of whole chromosomes and how they affect the phenotype of an organism and (2) changes in the structure of individual chromosomes and how they affect meiotic pairing. Human examples will be used to show the phenotypic consequences and methods for detection.

  • 9.E: Changes in Chromosome Number and Structure (Exercises)
  • 9.S: Changes in Chromosome Number and Structure (Summary)
  • 9.0: Changes in Chromosome Number
    If something goes wrong during cell division, an entire chromosome may be lost and the cell will lack all of these genes. The causes behind these chromosome abnormalites and the consequences they have for the cell and the organism is the subject of this section.
  • 9.1: Changes in Chromosome Structure
    If the chromosome is altered, but still retains the three critical features of a chromosome (centromeres, telomeres, and origin of replication), it will continue to be inherited during subsequent cell divisions, however the daughter cell may not retain all the genes. For example, if a segment of the chromosome has been lost, the cell may be missing some genes. The causes of chromosome structural abnormalites, which involves breaks in the DNA that makes up the chromosome.
  • 9.2: Chromosome Abnormalities in Humans
    To better understand the consequences let's consider those that affect people. As you will recall humans are 2n=46. The convention when describing a person's karyotype (chromosome composition) is to list the total number of chromosomes, then the sex chromosomes, and then anything out of the ordinary. Most of us are 46,XX or 46,XY. What follows are some examples of chromosome number and chromosome structure abnormalities.
  • 9.3: Diagnosing Human Chromosome Abnormalities
    How can we confirm that a person has a specific chromosomal abnormality? The first method was simply to obtain a sample of their cells, stain the chromosomes with Giemsa dye, and examine the results with a light microscope. Each chromosome can be recognized by its length, the location of its centromere, and the characteristic pattern of purple bands produced by the Giemsa.

Thumbnail: (Wikipedia-Pmx-CC:AS)


9: Changes in Chromosome Number and Structure - Biology

We previously learned how errors in mitosis can potentially lead to cancer. What could errors in meiosis result in? In this outcome, we’ll learn what happens when errors occur in chromosome number.

Learning Objectives

  • Identify a karyotype and describe its uses in biology
  • Identify common errors that can create an abnormal karyotype
  • Identify syndromes that result from a significant change in chromosome number

Structural Variations in Chromosomes | Genetics

The following point highlight the five main types of structural variation in chromosomes. The types are: 1. Deletion or Deficiency 2. Duplications 3. Translocations 4. Inversions 5. B-Chromosomes.

Type # 1. Deletion or Deficiency:

A deficiency means deletion of a small portion of a chromosome resulting in loss of one or more genes. A deficiency originates from breakage occurring at random in both chromatids of a chromosome (called chromosome break), or only in one chromatid (chromatid break).

The breakage may be caused by various agents such as radiation, chemicals, drugs or viruses at any time during the cell cycle, either in somatic or in germ cells. Depending upon its location, a deletion may be terminal when a single break occurs near the end of the chromosome or interstitial when two breaks occur in a middle portion of the chromosome.

Each break produces two raw ends which may behave in one of following three ways:

(a) There might be reunion of the broken ends called restitution so that the original chromosome structure is restored

(b) The broken ends may not unite giving rise to a chromosomal segment without a centromere which is eventually lost during cell division

(c) If two single breaks occur in two different chromosomes in a cell, the deleted segment of one chromosome may unite with the raw broken end on the other chromosome this is called exchange union.

Fate of a Deleted Fragment:

If the fragment does not have a centromere (acentric), then at metaphase it will not be able to get attached to spindle fibres and move towards a pole with other centric chromosomes. It will remain at the centre of the cell and will not be included within any of the two daughter nuclei. It will be free in the cytoplasm and will eventually be lost (Fig. 12.1). In this way, the cell will lose one or more genes contained in the deleted fragment.

A diploid cell has a homologue of the chromosome which has lost a segment. The corresponding segment of the intact homologue will have alleles of the genes that the cell has lost. Such a cell is said to be heterozygous for a deficiency. A very small deficiency in the heterozygous state is viable, but if homozygous it is lethal. When a deletion is large it is lethal even in the heterozygous state.

If a deletion occurs in cells of the germ line, then 50% of the gametes formed will have a deleted chromosome and 50% gametes would be normal. This would result in half the offspring with phenotypic abnormalities related to the genes carried on a small deleted fragment.

If the deficiency occurs in a developing embryo, some cells would have normal chromosomes and other cells would have the deficiency. This could produce a mosaic individual with two different phenotypes.

Detection of Deficiency:

The occurrence of a deficiency can sometimes be inferred from the results of a genetic cross when a rare recessive phenotype unexpectedly appears in the progeny. Consider a cross between two parents DD and dd where D controls the dominant expression of a trait, and d is the recessive allele. The F1 is expected to show the dominant trait and have the genotype Dd.

If on the contrary, some F1 individuals show the recessive phenotype, one explanation could be sought in a deletion of the chromosomal segment bearing gene D. Since other interpretations are also possible, it is best to confirm the occurrence of deficiency from a cytological study of the chromosomes as described below.

Deficiencies are best observed in preparations of homologously paired chromosomes at meiotic prophase either in large sized plant chromosomes or in polytene chromosomes. Normally during pachytene homologous chromosomes are intimately synapsed throughout their length.

If one of the homologues is deficient over a small length, the corresponding portion of the second homologue has nothing to pair with. It therefore, forms a loop (Fig. 12.1), which is clearly visible in cytological preparations and is clear-cut proof that deficiency has occurred.

Suppose further that the deleted segment carries the dominant gene D. The recessive allele d is still present in the other homologue and expresses itself even in the single dose because the dominant allele D which normally suppresses it is absent. Such a phenomenon whereby a single recessive allele expresses itself in absence of the dominant allele is called pseudo-dominance.

This also explains the results of the cross described above. Pseudo-dominance is similar to the hemizygous condition found exclusively in males where a recessive gene or a single X chromosome is expressed. Pseudo-dominance effect is observed in autosomal genes.

Due to the fact that deficiencies can produce unique phenotypic effects and the ease with which they can be identified by loop formation, they are important cytological tools for mapping genes.

In Drosophila this method has been used to locate a number of genes on the banded polytene chromosome. In general it can be stated that each band represents a distinct gene. Since there are 5000 bands in D. melanogaster it is believed that there are about 5,000 genes in this species.

Deficiencies also produce phenotypic abnormalities in man. The cat-cry syndrome (cri- du-chat) where the baby cries like a cat is due to a deletion in the short arm of chromosome 5. A deletion in the long arm of chromosome 22 (Ph 1 or Philadelphia chromosome) leads to chronic granulocytic leukemia.

In plants deficiencies are not easily transmitted to the progeny because their presence in developing pollen grains leads to pollen sterility. Nevertheless they have been observed in some plants.

In maize Creighton and McClintock have found that small deficiencies are viable even in the homozygous state. A special kind of single break occurs through the centromere of a metacentric chromosome giving rise to two isochromosomes with terminal telomeres. Such an event is also called misdivision of centromere (Fig. 12.2).

Type # 2. Duplications:

A duplication involves attachment of a chromosomal fragment resulting in addition of one or more genes to a chromosome. Whenever there is a duplication in a chromosome, there is a corresponding deletion in another chromosome.

Following types of duplications are known (Fig. 12.3: the diagrams are self-explanatory):

The phenotypic effect produced by a duplication is illustrated by the attached-X females in Drosophila. Consider such flies which are homozygous for some recessive sex-linked traits. It is found that when a fly receives a fragment of an X chromosome carrying the wild type allele from its male parent, then only the dominant phenotype is expressed.

The recessive alleles of the same gene although present in the homozygous condition, are not able to express themselves. Evidently the presence of a single dominant allele in a duplication is enough to produce the wild type phenotype.

The origin of duplications can be traced to unequal crossing over during meiosis. Normally homologous chromosomes are paired in a perfect manner so that identical loci lie exactly opposite each other.

The mechanism ensures that after crossing over between non-sister chromatids, equal exchange products are formed. If paired chromosomes are misaligned, it is not possible for exchange to take place between exactly opposite locations on two chromatids.

Instead, exchange occurs between adjacent points on two chromatids so that one resulting chromatid will have a duplication, the other a deletion. Such an exchange is called unequal crossing over. A gamete that receives a chromosome with a duplication will be diploid for some genes. When it fertilises a normal gamete, the zygote will have three sets of those genes that are present in the duplicated segment.

Bar eyes is a dominant X-linked trait in Drosophila females which provides a range of interesting phenotypes resulting from duplication. In a homozygous wild type female there is a large oval compound eye (non-bar) with about 779 facets.

The Bar trait reduces the eye to a vertical bar with very few facets. Bridges analysed the salivary gland chromosomes of Drosophila and found that the Bar gene (B) was present on a region designated 16A of the X chromosome.

When the band in the 16A region is present in duplicate in one X chromosome of the female (i.e. heterozygous for the duplication B/X), it results in an elongated Bar-shaped eye, smaller than the wild type (+/+) due to the presence of only 358 facets.

When a female is homozygous for the duplication (B/B), the Bar-shaped eye is further reduced in size and has 68 facets. If there is unequal crossing over in a female homozygous for Bar (B/B), it results in one chromatid where the 16A region (Bar locus) is present in triplicate, and the second chromatid with only one Bar locus.

Such a heterozygous triplicate condition produces a pheno­type known as ultra-bar (B u ) with only 45 facets. If the triplicate condition becomes homozygous (B U /B U ), the result is a very small eye with only 25 facets (Fig. 12.4). Unequal crossing over is also responsible for a rare human haemoglobin known as haptoglobin.

The Bar locus in Drosophila provides an explanation for position effect. According to this phenomenon the expression of a gene becomes altered when the position of the gene is physically changed. Cytologically, a duplication is identified by the same method as deficiency, since in the heterozygous condition the extra fragment forms a loop in one of the two homologues.

Type # 3. Translocations:

Sometimes a segment of a chromosome becomes detached and unites with another non-homologous chromosome. Such an inter-chromosomal rearrangement is called translocation.

The rearrangements are of following types (Fig. 12.5):

a. Simple Translocation:

A single break occurs in a chromosome, and the broken fragment becomes attached to the end of another chromosome. However, due to the presence of “non-sticky” telomeres at the unbroken ends of a chromosome, such a terminal attachment of a segment does not take place.

In this type three breaks are involved. Two breaks occur in a chromosome to produce an interstitial fragment. This fragment becomes inserted into one of the arms of another non-homologous chromosome in which a single break has produced two “sticky” ends.

c. Reciprocal Translocations:

These are the most frequent and extensively studied translocations. A single break occurs in each of the two non-homologous chromosomes followed by a mutual exchange of the broken fragments. This results in two new chromosomes each having one segment of the other chromosome.

Rarely, two breaks occur in each of the two chromosomes followed by exchange of intercalary segments. If the centromere containing segment of one chromosome is joined to the acentric piece of the other non-homologue, the exchange is called eucentric. But if two centric pieces from two non-homologues join to form a dicentric chromosome, it is called aneucentric.

In the next division the dicentric chromosome will form a bridge and the acentric fragment will be lost. Therefore aneucentric exchange unions are usually lethal. The eucentric reciprocal translocations produce viable gametes if both pairs of non-homologous chromosomes exchange segments.

Reciprocal exchange of segments involves no loss of genetic material. There is a qualitative change in the sequence of genes which is transmitted during mitosis and meiosis. Reciprocal translocations represent an important group of inter-chromosomal structural aberrations in chromosomes.

d. Multiple Translocations:

Sometimes more than two pairs of non-homologous chromosomes may be involved in a translocation as observed Drosophila and Oenothera. In 1930 Stern studied a multiple translocation system in Drosophila in which a segment of the Y chromosome became attached to the X chromosome. At the same time a reciprocal translocation occurred between the X and chromosome IV. This resulted in a female with 9 chromosomes instead of 8.

e. Half Translocations:

When the nucleus containing two broken chromosomes is small, the broken ends are not widely separated in space and have better chance of undergoing reciprocal exchange. This is true for the small compact nucleus in the head of a sperm.

In oocytes, on the contrary, due to the large nuclear volume the distance between the broken ends of non-homologous chromosomes may be so great that the chance for an exchange union is relatively small. In such a case only one exchange union may take place, leaving the other two broken ends free. This is called half translocation.

Cytology of Reciprocal Translocations:

In translocation homozygotes meiosis is normal with regular bivalent formation at pachytene. At anaphase, movement to the poles is normal and viable gametes are produced. On the contrary in translocation heterozygotes pairing is complicated due to segments that have been exchanged between non-homologous chromosomes.

Instead of bivalents therefore, cross-shaped configurations of quadrivalents are formed at Metaphase I because of synapsis between homologous segments. Such interchange figures are more easily recognised in plant species with large chromosomes.

A translocation heterozygote has two normal and two interchange chromosomes. Following the rules of pairing these four chromosomes will form a cross-shaped interchange configuration shown in Fig. 12.6. The fate of chromosomes and the type of gametes that will be produced depend upon the frequency and distribution of chiasmata.

The chance that one or more chiasmata are formed in a certain segment depends on three factors: the length of the segment, the amount of exchange in the given organism, and characteristic properties of the segment that relate to chiasma formation.

The formation of chiasma in the separate segments determines the frequencies of the different metaphase configurations. Conversely from the latter frequencies it is possible to estimate the chance that each segment has one or more chiasmata, or the intensity of crossing over in the segment.

When two opposite segments in the cross have a chiasma each, it results in two rod bivalents. If chaismata occur in two adjacent terminal segments, a trivalent and a univalent are formed. However, the ring and chain quadrivalents are the most common translocations.

Sometimes two heterozygous reciprocal translocations may occur in the same cell. When there is one chromosome common to both, a multivalent of chromosomes is formed (hexavalent). When two interchanges occur in the same two chromosomes a quadrivalent can result.

It is possible to determine whether or not two translocations share a chromosome. This is done by making a double heterozygote and by observing whether two quadrivalents, a hexavalent, or a single quadrivalent are formed. In plants like barley, Datura, maize, rye and some others, a series of interchanges are known, involving all chromosomes at least once.

The chromosomes involved in an interchange can be determined when the interchange is hybridised successively with all interchanges of the series, called a tester set. In one case a quadrivalent will be observed at meiosis the known interchange and the one to be analysed have both chromosomes in common.

In two cases a hexavalent is formed (the known and unknown interchange share one chromosome) and in the remaining cases two quadrivalents are observed. This is an efficient method of determining which chromosomes are involved in an unknown interchange.

In each of the four arms of the cross-shaped quadrivalent one chiasma is usually formed. At diakinesis two events take place: repulsion between homologues causing their separation, and terminalisation (movement) of chiasmata towards the distal end of the arms. At metaphase therefore, the interchange figure becomes oriented to form an open ring or a twisted, zigzag configuration (Fig. 12.7).

In case chiasma does not form in one of the four arms, the cross- shaped complex opens up to form a chain. Anaphasic movement of chromosomes towards the poles takes place in one of the three different ways described below (Fig. 12.8).

a. Alternate Segregation:

The twisted orientation ensures perfect disjunction so that both translocated chromosomes 1′ and 2′ go to one pole, and both un-translocated chromosomes 1 and 2 go to the other pole. Thus all the remaining gametes will receive a full complement of genes and would give rise to viable individuals.

b. Adjacent-1 Segregation:

This will take place in the open ring configuration whereby one translocated and one un-translocated chromosome will go to the same pole, in this way chromosome 1 and 2′ will go to one pole whereas 1’ and 2 will go to the other pole.

c. Adjacent-2 Segregation:

Again in the open ring configuration, two homologous chromosomes 1 and 1′ will go to one pole, the other two homologues 2 and 2′ go to the other pole. It is evident that both the adjacent types of segregation will give rise to gametes with duplications and deficiencies which would cause semi-sterility. The proportion of inviable gametes produced would be determined by the frequency of germ line cells having ring configuration.

In animals gametes with duplication deficiency genomes are viable, but the zygote does not survive. Homozygous translocations can give rise to viable individuals if the paired homologues have normal crossing over and segregation at meiosis.

The site in the cross-shaped figure where crossing over occurs is important in estimating sterility. With respect to crossing over, each arm has a distinct interstitial segment which lies between the centromere and the break point of the translocation the second is called pairing segment which represents portions of the arms of the cross beyond interstitial segments.

Crossing over in the pairing segments has no effect on the segregation pattern as only homologous are exchanged. The ring and zigzag arrangements obtained are in fact due to crossovers in the pairing segments. If crossing over occurs in the interstitial segment then non-homologous portions are exchanged leading to the production of unbalanced gametes.

Genetic Methods of Detecting Translocations:

Translocations can be detected by performing genetic crosses and observing gene segregation. When translocation heterozygotes are selfed or crossed with each other, the progeny is of three types: normal homozygotes, interchange heterozygotes, and interchange homozygotes in the ratio 1:2:1 (Fig. 12.9 A, B).

If there are two reciprocal translocations in a cell that do not share any chromosome, then there is independent segregation.

But if two translocations share a common chromosome there can be two consequences:

(a) The same homologue of the common chromosome is involved in both translocations. The resulting balanced gametes are of two types, one with both translocations, the other with none. Alternate segregation is necessary to produce them. Thus when heterozygotes are selfed the progeny is in the ratio: 1 homozygous for both, 2 heterozygous for both, and 1 homozygous normal.

(b) There is one chromosome shared by two translocations, but here one homologue is involved in one, the other homologue in the other translocation as indicated below.

This happens normally when two independently formed translocations are combined in one individual by hybridisation. Here also only two types of gametes are formed: one type having one translocation and the third chromosome normal the other having the other translocation and also one chromosome of the three normal.

When heterozygotes are now selfed they produce one homozygote for one translocation and also for one normal chromosome two double heterozygotes and the normal types are formed due to crossing over in the different segment.

Translocations affect linkage relationships between genes in two ways:

(a) In the homozygote linkage is changed the genes in the translocated segment are not linked with the genes in the chromosome where they originally belonged they are now linked to other genes. Study of this change in linkage can be used to detect a translocation and identify the involved chromosome.

(b) In the translocation heterozygote all the genes on all the involved chromosomes are linked. This is because usually only balanced gametes take part in fertilisation or only balanced zygotes survive. Balanced gametes arise when either all interchange chromosomes or all normal chromosomes are present in one gamete.

Recombination between genes on different chromosomes, that is, between the gene and translocation takes place between the interchange breakpoint and the locus. The percentage crossing over between a locus and the break point can be calculated. The translocation can be detected from mitotic chromosomes.

The heterozygotes can be located from multivalents at meiosis. The two homozygous types, the normal and interchange are not distinguishable from each other as both produce bivalents. An easier method of identifying heterozygotes is through their semi-sterility. The simplest analysis between the gene and translocation using heterozygotes can be done by a test cross which yields a 1: 1 segregation for the translocation and also for the gene.

Translocation in Oenothera:

The various species of the plant Oenothera (Onagraceae) are heterozygous for multiple translocations and show rings of chromosomes at meiosis. There are 14 chromosomes in the diploid cell of which some or all may be involved in translocations.

On this basis the species of Oenothera form a graded series. O. hookeri is distinct in having 7 pairs of chromosomes and no translocations. The other species form rings of 6, 8, 10, 12 or 14 chromosomes at meiosis (Fig. 12.10). O. lamarckiana has a ring of 12 chromosomes and one bivalent pair. In O. muricata all the 14 chromosomes are united to form a giant ring. O. biennis shows one ring of 8 and another of 6 chromosomes.

Similar instances of interchange heterozygosity are also known in some other plants such as Rhoeo discolor (Commelinaceae), Isotonia (Lobeliaceae), Hypericum (Hypericaceae) and 6 more genera of the family Onagraceae besides Oenothera. It is rare in animals.

A few genera of scorpions like Isometrus, Buthus and Tityus show translocation heterozygosity and ring of chromosomes at meiosis. There are certain genetic mechanisms which enforce permanent translocation heterozygosity in Oenothera. The cytogenetics of Oenothera has been worked out extensively.

Type # 4. Inversions:

Inversions result when there are two breaks in a chromosome and the detached segment becomes reinserted in the reversed order. They are classified into two types depending upon the inclusion or absence of the centromere within the inverted segment.

Thus when both breaks occur in one arm of the chromosome it leads to a paracentric inversion when a break occurs in each of the two arms, the centromere is included in the detached segment and leads to a pericentric inversion.

Meiosis is normal in inversion homozygotes. In heterozygotes pairing between homologous chromosomes is affected in the region of the inverted segment. Consequently, there is a suppression of recombination and fertility is impaired.

Paracentric Inversions:

This type of inversion is identified in the heterozygote by formation of a pairing loop at pachytene. If the size of the loop is large enough, chiasma formation will take place within it. When a single chiasma forms between an inverted and a normal segment, the two chromatids involved will produce one dicentric chromatid and one acentric fragment after exchange (Fig. 12.11). The other two chromosomes will be normal.

At anaphase I the dicentric chromosome will be pulled towards both poles, it will form a bridge that will ultimately break. The acentric fragment due to its inability to move would be eventually lost.

Consequently, of the resulting four gametes, two would be normal and two deficient in chromosome segments. In plants deficient gametes are not viable (pollen grains that are deficient usually abort and are nonfunctional). In animals such gametes take part in fertilisation but either the zygote or the embryo aborts.

In an individual heterozygous for a paracentric inversion therefore, viable offspring are produced only by two of the four chromatids which did not have chiasma formation between them in the region of the loop.

In each chromatid the gene sequence in the inversion segment will be of the non-recombinant, parental type. Consequently, none of the offspring would be recombinants for genes present within the inverted segment. In this way a paracentric inversion suppresses recombination throughout its length.

In some insects and in Drosophila, individuals heterozygous for an inversion do not show reduction in fertility. In fact paracentric inversions occur frequently in natural populations of Drosophila. There are two explanations for this. One is absence of crossing over in male meiosis.

The second is occurrence of four products of female meiosis in linear order of which the middle two egg nuclei have the deficiency the peripheral two nuclei are functional and fertilised. They produce viable offspring of the parental type.

Pericentric Inversions:

In an individual heterozygous for a pericentric inversion, the centromere is present within the loop. When chiasma formation takes place within the inverted segment the chromatids resulting after exchange do not form a dicentric and acentric fragment as in a paracentric inversion heterozygote.

Instead, they have one centromere each, but are deficient for some segments whereas other segments are duplicated (Fig. 12.12). The exchange segments produce inviable gametes and offspring. As in the case of pericentric inversions, the two chromatids not involved in crossing over only produce viable offspring with parental combination of genes present in the inverted segment.

Due to the suppression of recombination the genes present in the inverted segment segregate as a single unit called supergene within a population. Inversions are easy to identify in the banded polytene chromosomes of Drosophila larvae and have been extensively studied.

Type # 5. B-Chromosomes:

In addition to the normal chromosome complement, a number of plant and animal species have extra chromosomes called B-chromosomes (normal complement in such cases is designated A). They are smaller than the A chromosomes, they do not pair with any A chromosome during meiosis, and apparently do not serve any vital function in the organism.

However, they persist in the population without conferring any obvious advantages. Their number is variable within species and among individuals within a population, in others they may be lacking.

As they are not required for normal growth and reproduction, they have been considered to be genetically inert and dispensable. However, the recent work on corn and rye has shown that they have a few active genes and they perform certain functions.

B-chromosomes have been extensively studied in plants of Zea mays (maize) and Secale cereale (rye). They behave abnormally during mitotic division in the uninucleate pollen grains which gives rise to a small generative cell and a large vegetative cell. At anaphase of mitosis, the daughter chromatids of B-chromosomes fail to separate even though the centromeres have divided.

Due to nondisjunction both chromatids move together toward the pole which forms the generative nucleus. Later on when the generative nucleus divides to form two male gametes, B-chromosomes segregate normally. There is preferential fertilisation of eggs by male gametes which carry B-chromosomes.


Results

Identification of pairs of ohnologous genes in the ancestral Amniota genome

We inferred from Ensembl gene trees (version 69) that 19,786 genes existed in the ancestral Amniota genome, the ancestor of birds, reptiles, and mammals (326 million years). We used AGORA (Algorithm for Gene Order Reconstruction in Ancestors) [18] to order and orient these genes as they were in the Amniota genome (Additional file 1: Figure S1). This in silico reconstruction is composed of 470 segments, with 50% of the genes in segments larger than 253 genes (N50 length). We then selected the 56 chromosome-size segments larger than 50 genes as an initial set of Contiguous Ancestral Regions (CARs mean CAR length 256 genes, 12,134 genes total) to identify duplicated regions.

Ohnologs resulting from the 1R-2R WGDs are key to identifying pairs of duplicated chromosome segments. We identified pairs of putative ohnologous genes directly in the reconstructed Amniota genome using gene trees to date their duplication while ensuring that each member of a pair belongs to a different CAR (see the “Methods” section), resulting in a “List A” containing 5616 ancestral Amniota ohnolog pairs. Two previous studies have also identified ohnologs from the two WGDs, in the human and other vertebrate genomes, using conserved synteny and sequence similarity. We used Ensembl gene trees to convert extant gene identifiers from these studies to their ancestral Amniota gene identifiers. The first list established by Makino and McLysaght [8] and hereafter called “List B” contains 4870 ancestral Amniota gene pairs. The second study by Singh et al. [9] established three levels of confidence (strict, intermediate, and relaxed) to define ohnologs. Following these criteria, we defined three additional lists containing respectively 2873 (List “C-strict”), 5253 (List “C-inter”), and 7806 (List “C-relax”) ancestral Amniota ohnolog pairs.

The sum of the A, B, and C-relax lists shows only 25% of genes in common (Fig. 2a), but we show below that the three lists nevertheless support the 1R-2R hypothesis. In this scenario, each original chromosome is duplicated in two then four copies chromosomes thus form tetrads where each possesses three ohnologous counterparts. We tested whether pairs of CARs share more ohnologs than expected if they were distributed randomly using each of the five lists of ohnologs (proportionality test see the “Methods” section). We show that, in all cases, CARs are ohnologous to three other CARS on average (Additional file 1: Figure S2). Despite their differences, all lists therefore support the 1R-2R hypothesis, justifying the construction of an improved consensus list of ancestral Amniota ohnolog pairs using all five lists. We started from the intersection of lists A, B, and C-strict as the most reliable subset (1273 pairs of ohnologs) and gradually extended it by adding pairs of genes from lower confidence subsets (pairs of genes intersecting fewer lists, or lists that include C-inter and C-relaxed Fig. 2b). In this process, we ensured that the growing list remained compatible with the 1R-2R hypothesis: a pair could never be included if the two genes belong to two different phylogenetic trees (i.e., the two duplicated genes in a pair must descend from a common ancestral gene) and all the ohnologs within a phylogenetic tree, when arranged in pairs, cannot link more than four CARs (a tetrad Fig. 2b and Additional file 1). This incremental process (Additional file 2: Table S5) resulted in a list of 8184 ohnologous genes, forming 7441 ohnolog pairs grouped into 2973 ohnolog families, each family in principle corresponding to one pre-1R gene (Additional file 3, 4, and 5 respectively for the list of ohnolog genes along with their human descendants, the list of ohnolog pairs and the list of ohnolog families).

Identification of ohnolog pairs in the ancestral Amniota genome. (a) Comparison between five lists of ohnolog pairs in Amniota. Left: a Venn diagram of the sets of ohnolog pairs from five lists: list A (this study), list B [8] and the three lists C [9]. The numbers of pairs at the intersections of the lists are indicated. Right: a Venn diagram of the sets of ohnolog genes from the same lists as above. The overlap between the lists of ohnolog genes is higher than between the lists of pairs because the latter contain different pairs between the same genes. For example, two pairs G1-G2 and G1-G4 are in different lists (no overlap between lists) but gene G1 is common to both lists (1 gene overlap see (b) for a graphical illustration). The surface of the circles and their intersection are roughly proportional to the number of genes pairs or genes of each list. (b) Schematic example of ohnolog pair selection. Step 1: from the initial list of 1273 gene pairs (black area in Venn diagram), 2 pairs involve 3 genes G1, G2, and G3, each on a different CAR. Step 2: pairs from a new sub-list are considered, a new gene pair G1-G4 is added to the network. Gene G4 is on a fourth CAR. Step 3: A new list is considered, a new pair is identified (G4-G5) but G5 is on a fifth CAR so pair G4-G5 is discarded. Step 4: a new list is considered, a pair G4-G3 supporting the network is identified

This list of ohnolog pairs is of high quality for a number of reasons. First, it is based on the gene content and synteny in the reconstructed ancestral Amniota genome which is 326 million years closer to the 1R-2R events than extant genomes, and thus, signatures of 1R-2R events are read with greater accuracy. Second, this list abides by a 1R-2R-compatibility rule, i.e., no ohnologous gene family connects more than four CARs. Third, the ohnologous pairs are all phylogenetically consistent in that both genes in a pair always belong to the same Ensembl gene tree. Fourth, the two genes of a pair were allowed to be on the same CAR only if ≥ 90 genes separated them to avoid spurious inclusions of genes duplicated in tandem.

Identification of post-2R duplicated CARs

Using our improved list of ohnolog pairs, we manually split, assembled, and grouped ancestral Amniota CARs in order to convert them to a configuration that is as close as possible to the post-2R karyotype. In the simplest scenario, post-2R CARs should readily form tetrads of four ohnologous CARs, each corresponding to one pre-1R chromosome. However, chromosome rearrangements between the 1R and 2R, between the 2R and Amniota, and incomplete or incorrect reconstruction of CARs all concur to disrupting this ideal pattern. We started with the 56 largest CARs and applied the proportion test to identify CARs sharing a significant number of ohnologous genes as described above (i.e., ohnologous CARs). We identified groups of at least three CARs all significantly ohnologous pairwise (p value < 5.10 −2 , Bonferroni adjusted). These were completed into tetrads (i.e., four CARs all significantly ohnologous to each other) by including smaller CARs and/or CARs at lower significance thresholds. We also merged CARs that showed evidence of belonging to the same Amniota chromosome (Table 1), because they were merged in alternative AGORA reconstructions using different sets of parameters, and/or they showed identical homologies to Amniota descendent genomes (human or chicken Fig. 1). In addition, merged CARs had to be significantly ohnologous to at least one CAR in common in a triad and show no significant ohnology with each other. We also split CARs that showed, along their length, a disruption in their distribution of ohnologs and disruption of their homologies to chicken, human, spotted gar, or medaka chromosomes (Fig. 1). Finally, we confirmed merged CARs using homologies with outgroup species such as the spotted gar or the medaka (Fig. 1 see the “Methods” section).

The resolution of CARs in tetrads is much clearer after this conversion of Amniota CARs to a post-2R configuration, especially compared to human chromosomes (Fig. 3a). The proportionality test links each curated Amniota CAR on average to 3 other CARs almost independently of any p value threshold, whereas human chromosomes are much more sensitive to the p value threshold. Indeed in the human genome, the expected average of three partners per chromosome is reached only at a p value of 1.10 −09 (Fig. 3b), a stringent threshold where 7 human chromosomes cannot be assigned to a tetrad. An example of construction of a tetrad and assembly of CARs is detailed in Fig. 3c. We identified three significantly ohnologous CARs grouped in a triad (CARs 73, 117, 250). Two additional CARs (256 and 137) show significant ohnology to CAR 250. The tetrad was completed with the addition of a smaller CAR (CAR 82), which was linked to four of the five initial CARs with significant but higher p values (Additional file 6). Then, of the five initial CARs, three fulfilled the conditions to be assembled in a single larger CAR (CARs 117, 137, 256): they were ohnologous to CARs in common but were not ohnologous to each other, and AGORA merged CARs 117/137 and CARs 137/256 together when using a more recent version of Ensembl (Version 84). Furthermore, the three CARs map to the same chicken and spotted gar chromosomes, strongly suggesting that they derive from the same chromosome of their common Vertebrata ancestor. Finally, all three assembled CARs, when mapped on the medaka genome (a teleost fish that went through an additional WGD [19]), are orthologous to the same two medaka chromosomes (13 and 14 Additional file 6).

Organization of ancestral Amniota CARs in tetrads (a) Circos plot [45] showing the pairs of ohnologs involving each of the four chromosomes (Homo sapiens) or CARs (Amniota) of the tetrad carrying the Hox genes (Tetrad 1 in D). The pairs of ohnologs in the human genome were the descendants of those of Amniota (6121 pairs of human ohnologs vs. 7441 pairs of amniote ohnologs). The human Hox cluster tetrad is mainly composed of human chromosomes 2, 7, 12, and 17. The Amniota Hox cluster tetrad is composed of CARs 108, 24, 99, and 6_39_140. An ohnolog pair is represented (green lines) between two Amniota CARs or two human chromosomes if at least one of the two genes of the pair falls on a chromosome/CAR of the tetrad. The Amniota Hox CARs are involved in 634 pairs, while the human Hox chromosomes are involved in 2171 pairs of ohnologs. This figure shows that the reconstruction of Amniota ancestor displays a clearer picture of the 1R-2R than the human genome. (b) Ohnolog partners per CAR/chromosome in the Amniota (left) and human (right) genomes. Each boxplot shows the distribution of the number of CARs (Amniota) or chromosomes (Human) found to be ohnologous to a given CAR/chromosome by the proportionality test. The x-axis shows the Bonferroni adjusted p value thresholds used to select ohnologous chromosome/CARs. Triangles indicate the average number of partners. The Amniota genome shows a clear and stable distribution of three partners per CAR across a wide range of p values, as expected after two WGDs where chromosomes are grouped in tetrads. In contrast, the distribution in Homo sapiens shows that extremely low p value thresholds must be used to reach the expected average of three partners, justifying the fragmentation of the human genome as described in [6]. (c) Example of how a group of significantly ohnologous CARs was analyzed to form tetrad 3. Black double-headed arrows (p value < 5.10 −2 after Bonferroni adjustment) represent the raw output of the proportion test, showing CARs with significant ohnology relationships. CARs 73, 117, and 250 form a triad of mutually ohnologous CARs. Dotted lines are additional ohnologous relationships that are supported without the Bonferroni adjustment. Numbers in black indicate CARs of at least 50 genes, while smaller CARs (< 50 genes) are in grey. Additional evidence (see text) was used to complete the tetrad. (d) Seventeen tetrads composed of 51 CARs. CARs are numbered arbitrarily and are joined by underscores in an arbitrary order when assembled. The letters “a” or “b” indicate that the CAR has been split in two segments (CARs 5 and 118) as part of the conversion to a post-2R karyotype (see text) and one CAR is present twice in two different tetrads (CAR 10_240_2) to facilitate the representation (pale yellow shapes)

We identified two post-2R chromosomal fusions that required a split of two Amniota CARs in two sub-CARs each (CARs 5 and 118), and we identified three probable assembly errors that required a split of three CARs (CARs 40, 46, and 97). We also performed 23 CAR assemblies (Table 1), ending with a final set of 51 CARs edited to more closely represent their post-2R configuration. The ohnology relationships between CARs based on significant p values of the proportion test connected the 51 CARs into 17 tetrads (Fig. 3d). The precise step-by-step procedure that we followed to split or assemble CARs and group them in tetrads is detailed in Additional file 1, including Additional file 1: Figure S7B to S14B.

Chromosome evolution between the 1R and 2R whole genome duplications

The 17 tetrads composed of 51 Amniota CARs are not all disjoint: some share one or two CARs in common, reflecting chromosomal events between the 1R and the 2R, and after the 2R. We identified these events by first establishing theoretical scenarios corresponding to each configuration. A single disjoint tetrad implies a simple evolutionary scenario without any large chromosomal rearrangement between the two WGDs (Fig. 4a). Two adjacent tetrads, however, can be explained by one of two scenarios, each with the same degree of parsimony: a post-1R chromosome descending from a single pre-1R chromosome was broken (a fission), or two post-1R chromosomes, each descending from separate pre-1R chromosomes, were merged (a fusion Fig. 4b). As previously noted [20, 21], a non-duplicated outgroup species would be helpful to discriminate between the two possible ancestral configurations. However, of the two nearest outgroups to vertebrates (Fig. 1), neither tunicates (e.g., species of the Ciona group) nor cephalochordates (e.g., the amphioxus Branchiostoma floridae) [7, 22] are suitable for this purpose. The former are too diverged to identify clear chromosome homologies, and the genome of B. floridae is too fragmented to be informative. To circumvent this problem, we used a previously published reconstruction of 17 Chordate Linkage Groups (CLG) (Additional file 1: Figure S15), which are groups of human genes descended from the same ancestral chordate chromosome [7]. This reconstructed proto-karyotype precedes the 1R-2R events by less than 50 million years and is located at a much shorter evolutionary distance to the Vertebrata ancestor than the extant amphioxus genome (Fig. 1).

Evolutionary scenario models. (a) A single evolutionary scenario explains the formation of a single disjoint tetrad of ohnologous CARs. (b) Two equally possible evolutionary scenarios can explain how ohnologous CARs can form two adjacent tetrads: a fission or a fusion of chromosomes could have occurred between the 2 WGDs. In each case, the B and D chromosomes each possess two distinct parts (dark and light grey) homologous to distinct chromosome sets. B and D are therefore common to two tetrads. (c) A chromosome fusion after the two WGDs explains how two tetrads can be joined via a single CAR

Remarkably, each CLG was associated with one predominant CAR tetrad (Additional file 1: Figure S16). Consequently, all seven adjacent tetrads result from chromosome fusions between the 1R and 2R WGDs. Indeed, a chromosome fission would have split the gene content of a CLG over two different tetrads (Fig. 4b). Incidentally, we also found evidence for a chromosome fusion between the chordate ancestor and the 1R, because more than 75% of genes from CLGs 6 and 7 have their descendants in a single tetrad (tetrad 14 Additional file 1: Figure S16 and Additional file 7). Conversely, we could not confidently assign tetrad 13 to a CLG, likely because it is a small tetrad. We therefore conclude that the pre-1R karyotype comprised 17 chromosomes, duplicated into 34 chromosomes after the first WGD and followed by seven fusions. The resulting 27 chromosomes were duplicated in the second WGD leading to 54 Vertebrata chromosomes, at the origin of the approximately 60,000 extant species of vertebrates.

Chromosome evolution after the 2R

This karyotype was followed by additional chromosome fusions at different stages after the 2R WGD. Four fusions followed the scenario described in Fig. 4c, where a single CAR joins two tetrads (CAR 5, CAR5a_152, CAR3_22, CAR 10_240_2 Fig. 3d). They can be dated to the period between the 2R WGD and the Euteleostomi ancestor because of their homologies to both descendent and outgroup genomes. For example, CAR_10_240_2 is homologous to a single chicken chromosome (GG4), a single human chromosome (chromosome X) and a single spotted gar chromosome (LG7), which is most parsimoniously consistent with a situation where this CAR was already a single chromosome in Euteleostomi, the common ancestor of these three species. A fifth fusion can be dated to the period between Euteleostomi and Amniota: CAR 118 is common to two tetrads but while it is homologous to a single chicken chromosome (GG1), a disruption of synteny in the spotted gar genome and a disruption in the DCS pattern in the medaka genome are consistent with a fusion in the lineage leading to Amniota.

Accounting for these fusions, the 54 chromosomes in the post-2R Vertebrata led to a Eueteleostomi karyotype of 50 chromosomes (4 fusions) and to an Amniota karyotype of 49 chromosomes (1 fusion Fig. 5 and Additional file 1). A dedicated Genomicus server [23] provides a graphical interface to compare and analyze the genomes presented in this work (Additional file 1: Figure S17 http://genomicus.biologie.ens.fr/genomicus-69.10/).

Reconstructed evolutionary history of karyotypes from Chordata to Amniota. On the right, a simplified species tree of the Chordata is shown, with WGD events depicted by red stars. The eight lineages represented from left to right are mammals, birds, teleost fish, holostocean fish (gar), cartilaginous fish, cyclostomes (lamprey, hagfish), tunicates (ciona), and cephalochordates (amphioxus). On the left, successive reconstructed karyotypes are shown, with one color for each of the 17 pre-1R chromosomes. The length of each pre-1R chromosome is proportional to its number of genes. For the 17 Chordate Linkage Groups (CLGs) of [7], the size of the colored segment is proportional to the number of genes that are found in the intersection of the CLG with a pre-1R chromosome, although segments corresponding to < 10% of the number of genes of the CLG were omitted for clarity (Additional file 1: Table S7). The karyotype between 1R and 2R was deduced from the pre-1R karyotype and the seven chromosome fusions are shown with purple curvy lines joining the fused chromosomes. The Euteleostomi karyotype was deduced from the Vertebrata karyotype after four chromosome fusions (Additional file 1). The lengths of the Euteleostomi chromosomes are proportional to the number of genes in the homologous Amniota CARs. Finally, the Amniota karyotype differs from that of Euteleostomi by only one chromosome fusion. The Amniota chromosomes were numbered from 1 to 49 (Additional file 1: Table S11 for correspondence with the CARs and number of genes). Black stars under 12 Euteleostomi chromosomes denote predicted ancestral micro-chromosomes

Comparative genomics between the pre-1R genome and the human genome

To enable comparisons between the pre-1R vertebrate ancestor genome reconstruction and extant species, we assigned genes to each of the 17 pre-1R chromosomes. Among Amniota genes, only ohnologs could be confidently assigned to pre-1R chromosomes, as non-ohnolog genes could have been acquired after the 1R WGD. To circumvent this problem, we applied a conservative procedure to assign 5052 of the 10,093 ancestral Olfactores genes to the 17 predicted pre-1R chromosomes (Additional file 8). The Olfactores ancestor is the common ancestor of vertebrates and tunicates and the closest ancestor upstream of the reconstructed pre-1R genome in Ensembl gene trees. This set of ancestral genes provides a direct connection to the human genome, through their 8378 human descendent genes. By identifying each of these human descendants by the color of its ancestral pre-1R chromosome (Fig. 6), we show that the structure of the 17 pre-1R chromosomes is still strikingly apparent in the human genome, with some chromosomes almost entirely composed of genes from a single pre-1R chromosome (e.g., chromosomes 14 and 15). We measured the degree of conservation of the post-1R-2R ohnolog content in windows of 50 genes positioned every 10 genes across human chromosomes. Three regions overlapping the Hox clusters A, B, and D stand out (and Hox C to a lesser degree Fig. 6), in line with the known functional importance associated with the clustering of these ohnologs [24].

Comparison between the pre-1R karyotype (top) composed of 17 chromosomes and the human karyotype (middle). The 8282 known human descendent genes of pre-1R genes are drawn at their position in the human genome with the color of their pre-1R ancestral chromosome. The position of 12 extant clusters (4 HOX, 4 FOX and 4 MHC) descending from a single clusters in pre-1R chromosomes are indicated by a black circle and a 2-character identifier (M1, M2, M3, M4 for MHC clusters, F1, F2, F3, F4 for FOX clusters, HA, HB, HC, HD for HOX clusters). A second human karyotype (bottom) shows, in a white-to-red scale, the number of ohnologs in windows of 50 genes positioned every 10 genes. Open circles denote the position of HOX clusters. Human chromosomes are drawn to scale, in Mb. Pre-1R chromosomes are drawn as in Fig. 5, in proportion to the number of genes assigned to each. The order of genes in pre-1R chromosomes being unknown, the positions of the 3 pre-1R gene clusters within their chromosome are arbitrary

The four Hox clusters originate from pre-1R chromosome 1, and we examined in the same light other paralogous clusters that have been proposed to originate from a single pre-1R locus. The MHC region on human chromosome 6 contains a number of genes unrelated to immune functions but which possess ohnologs in 3 other loci on chromosomes 1, 9, and 19 [25]. All 4 regions descend from pre-1R chromosome 9. Similarly, the loci containing FOX gene clusters have been compared and found to share paralogs suggestive of en-bloc duplications early in vertebrate evolution [26]. We confirm here their unique origin on pre-1R chromosome 10. In contrast, no common origin was found for the different clusters of imprinted genes in the human genome (e.g., H19 locus on chromosome 11, IGF2R locus on chromosome 6, PON [1,2,3] locus on chromosome 7, UBE3A locus on chromosome 15) [27, 28], in line with their known progressive appearance later in therian mammals [29]. The evolutionary scenario of ancestral vertebrate chromosomes presented here is therefore consistent with our current view of the evolution of these important gene families.

Finally, we analyzed the frequency of Gene Ontology terms of the human descendants of the 1416 pairs (2 gene losses), 502 triplets (1 gene loss), and 172 quartets (no loss) of ancestral Amniota ohnologs and find a striking pattern: quartets are enriched in both neuronal development and neuronal function (synaptic transmission) and triplets are enriched in muscle development (especially heart) and in muscle function (contraction), while pairs (two losses) are enriched in protein maturation and transport between organelles (Additional file 9).


AQA AS Biology Paper 2 | Tuesday 6 June 2017

(i)
The student put the same volume of water in each tube.
Explain why it was important that he controlled this experimental variable.

2. (membrane) proteins denature

2. (Only) one of the strands/template strand is used (to make mRNA/is transcribed)

3. (Complementary) base pairing so A→U, T→A, C→G, G→C

4. (RNA) nucleotides joined by RNA polymerase

(i)
What is the effect of Patau syndrome on the chromosomes of this female?

(ii)
Describe how the change in chromosome number in Patau syndrome was produced.

(ii)
1. In meiosis
2. Homologous chromosomes / sister chromatids do not separate

(iii)
1. Mutation / extra chromosome in gamete/egg/sperm (that formed zygote)

2. All cells derived (from a single cell/zygote) by mitosis
OR
3. All cells derived from a single cell/zygote by mitosis


DMCA Complaint

If you believe that content available by means of the Website (as defined in our Terms of Service) infringes one or more of your copyrights, please notify us by providing a written notice (“Infringement Notice”) containing the information described below to the designated agent listed below. If Varsity Tutors takes action in response to an Infringement Notice, it will make a good faith attempt to contact the party that made such content available by means of the most recent email address, if any, provided by such party to Varsity Tutors.

Your Infringement Notice may be forwarded to the party that made the content available or to third parties such as ChillingEffects.org.

Please be advised that you will be liable for damages (including costs and attorneys’ fees) if you materially misrepresent that a product or activity is infringing your copyrights. Thus, if you are not sure content located on or linked-to by the Website infringes your copyright, you should consider first contacting an attorney.

Please follow these steps to file a notice:

You must include the following:

A physical or electronic signature of the copyright owner or a person authorized to act on their behalf An identification of the copyright claimed to have been infringed A description of the nature and exact location of the content that you claim to infringe your copyright, in sufficient detail to permit Varsity Tutors to find and positively identify that content for example we require a link to the specific question (not just the name of the question) that contains the content and a description of which specific portion of the question – an image, a link, the text, etc – your complaint refers to Your name, address, telephone number and email address and A statement by you: (a) that you believe in good faith that the use of the content that you claim to infringe your copyright is not authorized by law, or by the copyright owner or such owner’s agent (b) that all of the information contained in your Infringement Notice is accurate, and (c) under penalty of perjury, that you are either the copyright owner or a person authorized to act on their behalf.

Send your complaint to our designated agent at:

Charles Cohn Varsity Tutors LLC
101 S. Hanley Rd, Suite 300
St. Louis, MO 63105


Unless otherwise stated, custom code used in this study is available at https://github.com/abmudd/Assembly.

Wurster, D. H. & Benirschke, K. Indian muntjac, Muntiacus muntjak: a deer with a low diploid chromosome number. Science 168, 1364–1366 (1970).

Wurster, D. H. & Benirschke, K. Chromosome studies in some deer, the springbok, and the pronghorn, with notes on placentation in deer. Cytologia 32, 273–285 (1967).

Chi, J. et al. New insights into the karyotypic relationships of Chinese muntjac (Muntiacus reevesi), forest musk deer (Moschus berezovskii) and gayal (Bos frontalis). Cytogenet. Genome Res. 108, 310–316 (2005).

Hsu, T. C., Pathak, S. & Chen, T. R. The possibility of latent centromeres and a proposed nomenclature system for total chromosome and whole arm translocations. Cytogenet. Cell Genet. 15, 41–49 (1975).

Liming, S., Yingying, Y. & Xingsheng, D. Comparative cytogenetic studies on the red muntjac, Chinese muntjac, and their F1 hybrids. Cytogenet. Genome Res. 26, 22–27 (1980).

Lin, C. C., Sasi, R., Fan, Y.-S. & Chen, Z.-Q. New evidence for tandem chromosome fusions in the karyotypic evolution of Asian muntjacs. Chromosoma 101, 19–24 (1991).

Scherthan, H. Localization of the repetitive telomeric sequence (TTAGGG)n in two muntjac species and implications for their karyotypic evolution. Cytogenet. Cell Genet. 53, 115–117 (1990).

Lee, C., Sasi, R. & Lin, C. C. Interstitial localization of telomeric DNA sequences in the Indian muntjac chromosomes: further evidence for tandem chromosome fusions in the karyotypic evolution of the Asian muntjacs. Cytogenet. Genome Res. 63, 156–159 (1993).

Yang, F., Carter, N. P., Shi, L. & Ferguson-Smith, M. A. A comparative study of karyotypes of muntjacs by chromosome painting. Chromosoma 103, 642–652 (1995).

Frönicke, L., Chowdhary, B. P. & Scherthan, H. Segmental homology among cattle (Bos taurus), Indian muntjac (Muntiacus muntjak vaginalis), and Chinese muntjac (M. reevesi) karyotypes. Cytogenet. Genome Res. 77, 223–227 (1997).

Yang, F., O’Brien, P. C. M., Wienberg, J. & Ferguson-Smith, M. A. A reappraisal of the tandem fusion theory of karyotype evolution in the Indian muntjac using chromosome painting. Chromosome Res. 5, 109–117 (1997).

Wang, W. & Lan, H. Rapid and parallel chromosomal number reductions in muntjac deer inferred from mitochondrial DNA phylogeny. Mol. Biol. Evol. 17, 1326–1333 (2000).

Chi, J. X. et al. Defining the orientation of the tandem fusions that occurred during the evolution of Indian muntjac chromosomes by BAC mapping. Chromosoma 114, 167–172 (2005).

Hartmann, N. & Scherthan, H. Characterization of ancestral chromosome fusion points in the Indian muntjac deer. Chromosoma 112, 213–220 (2004).

Zhou, Q. et al. Comparative genomic analysis links karyotypic evolution with genomic evolution in the Indian muntjac (Muntiacus muntjak vaginalis). Chromosoma 115, 427–436 (2006).

Tsipouri, V. et al. Comparative sequence analyses reveal sites of ancestral chromosomal fusions in the Indian muntjac genome. Genome Biol. 9, R155 (2008).

Farré, M. et al. Evolution of gene regulation in ruminants differs between evolutionary breakpoint regions and homologous synteny blocks. Genome Res. 29, 576–589 (2019).

Chen, L. et al. Large-scale ruminant genome sequencing provides insights into their evolution and distinct traits. Science 364, eaav6202 (2019).

Zimin, A. V. et al. A whole-genome assembly of the domestic cow, Bos taurus. Genome Biol. 10, R42 (2009).

Bana, N. Á. et al. The red deer Cervus elaphus genome CerEla1.0: sequencing, annotating, genes, and chromosomes. Mol. Genet. Genomics 293, 665–684 (2018).

Li, Z. et al. Draft genome of the reindeer (Rangifer tarandus). Gigascience 6, 1–5 (2017).

Stephens, P. J. et al. Massive genomic rearrangement acquired in a single catastrophic event during cancer development. Cell 144, 27–40 (2011).

Weisenfeld, N. I., Kumar, V., Shah, P., Church, D. M. & Jaffe, D. B. Direct determination of diploid genome sequences. Genome Res. 27, 757–767 (2017).

Lieberman-Aiden, E. et al. Comprehensive mapping of long-range interactions reveals folding principles of the human genome. Science 326, 289–293 (2009).

Lin, C.-C. et al. Construction of an Indian muntjac BAC library and production of the most highly dense FISH map of the species. Zool. Stud. 47, 282–292 (2008).

Jiang, Y. et al. The sheep genome illuminates biology of the rumen and lipid metabolism. Science 344, 1168–1173 (2014).

Schneider, V. A. et al. Evaluation of GRCh38 and de novo haploid genome assemblies demonstrates the enduring quality of the reference assembly. Genome Res 27, 849–864 (2017).

Jones, P. et al. InterProScan 5: genome-scale protein function classification. Bioinformatics 30, 1236–1240 (2014).

Slate, J. et al. A deer (subfamily Cervinae) genetic linkage map and the evolution of ruminant genomes. Genetics 160, 1587–1597 (2002).

Frohlich, J. et al. Karyotype relationships among selected deer species and cattle revealed by bovine FISH probes. PLoS ONE 12, e0187559 (2017).

Huang, L. et al. High-density comparative BAC mapping in the black muntjac (Muntiacus crinifrons): molecular cytogenetic dissection of the origin of MCR 1p+4 in the X1X2Y1Y2Y3 sex chromosome system. Genomics 87, 608–615 (2006).

Huang, L., Wang, J., Nie, W., Su, W. & Yang, F. Tandem chromosome fusions in karyotypic evolution of Muntiacus: evidence from M. feae and M. gongshanensis. Chromosome Res. 14, 637–647 (2006).

Toljagić, O., Voje, K. L., Matschiner, M., Liow, L. H. & Hansen, T. F. Millions of years behind: slow adaptation of ruminants to grasslands. Syst. Biol. 67, 145–157 (2018).

Zurano, J. P. et al. Cetartiodactyla: updating a time-calibrated molecular phylogeny. Mol. Phylogenet. Evol. 133, 256–262 (2019).

Ma, S., Wang, Y. & Xu, L. Taxonomic and phylogenetic studies on the genus Muntiacus. Acta Theriol. Sin. 6, 190–209 (1986).

Dong, W., Pan, Y. & Liu, J. The earliest Muntiacus (Artiodactyla, Mammalia) from the Late Miocene of Yuanmou, southwestern China. C. R. Palevol 3, 379–386 (2004).

Kumar, S., Stecher, G., Suleski, M. & Hedges, S. B. TimeTree: a resource for timelines, timetrees, and divergence times. Mol. Biol. Evol. 34, 1812–1819 (2017).

Maruyama, T. & Imai, H. T. Evolutionary rate of the mammalian karyotype. J. Theor. Biol. 90, 111–121 (1981).

Bush, G. L., Case, S. M., Wilson, A. C. & Patton, J. L. Rapid speciation and chromosomal evolution in mammals. Proc. Natl Acad. Sci. USA 74, 3942–3946 (1977).

The Chimpanzee Sequencing and Analysis Consortium. Initial sequence of the chimpanzee genome and comparison with the human genome. Nature 437, 69–87 (2005).

Locke, D. P. et al. Comparative and demographic analysis of orang-utan genomes. Nature 469, 529–533 (2011).

IJdo, J. W., Baldini, A., Ward, D. C., Reeders, S. T. & Wells, R. A. Origin of human chromosome 2: an ancestral telomere-telomere fusion. Proc. Natl Acad. Sci. USA 88, 9051–9055 (1991).

Huang, L., Chi, J., Nie, W., Wang, J. & Yang, F. Phylogenomics of several deer species revealed by comparative chromosome painting with Chinese muntjac paints. Genetica 127, 25–33 (2006).

Zou, Y., Yi, X., Wright, W. E. & Shay, J. W. Human telomerase can immortalize Indian muntjac cells. Exp. Cell Res. 281, 63–76 (2002).

Liming, S. & Pathak, S. Gametogenesis in a male Indian muntjac x Chinese muntjac hybrid. Cytogenet. Genome Res. 30, 152–156 (1981).

Ghavi-Helm, Y. et al. Highly rearranged chromosomes reveal uncoupling between genome topology and gene expression. Nat. Genet. 51, 1272–1282 (2019).

Baker, R. J. & Bickham, J. W. Karyotypic evolution in bats: evidence of extensive and conservative chromosomal evolution in closely related taxa. Syst. Biol. 29, 239–253 (1980).

Marks, J. Rates of karyotype evolution. Syst. Zool. 32, 207–209 (1983).

Gladkikh, O. L. et al. Rapid karyotype evolution in Lasiopodomys involved at least two autosome—sex chromosome translocations. PLoS ONE 11, e0167653 (2016).

Nash, W. G., Wienberg, J., Ferguson-Smith, M. A., Menninger, J. C. & O’Brien, S. J. Comparative genomics: tracking chromosome evolution in the family Ursidae using reciprocal chromosome painting. Cytogenet. Genome Res. 83, 182–192 (1998).

Carbone, L. et al. Gibbon genome and the fast karyotype evolution of small apes. Nature 513, 195–201 (2014).

Gordon, D. J., Resio, B. & Pellman, D. Causes and consequences of aneuploidy in cancer. Nat. Rev. Genet. 13, 189–203 (2012).

Funk, W. C., Zamudio, K. R. & Crawford, A. J. Advancing understanding of amphibian evolution, ecology, behavior, and conservation with massively parallel sequencing. in Population Genomics (eds Hohenlohe, P. & Rajora, O. P.), https://doi.org/10.1007/13836_2018_61 (Springer, 2018).

De La Torre, A. R. et al. Insights into conifer giga-genomes. Plant Physiol. 166, 1724–1732 (2014).

Drpic, D. et al. Chromosome segregation is biased by kinetochore size. Curr. Biol. 28, 1344–1356 (2018). e5.

Hockemeyer, D., Sfeir, A. J., Shay, J. W., Wright, W. E. & de Lange, T. POT1 protects telomeres from a transient DNA damage response and determines how human chromosomes end. EMBO J. 24, 2667–2678 (2005).

Camacho, C. et al. BLAST+: architecture and applications. BMC Bioinform. 10, 421 (2009).

Shi, Y. F., Shan, X. N., Li, J., Zhang, X. M. & Zhang, H. J. Sequence and organization of the complete mitochondrial genome of the Indian muntjac (Muntiacus muntjak). Acta Zool. Sin. 49, 629–636 (2003).

Zhang, X. M. et al. Muntiacus reevesi mitochondrion, complete genome. NCBI Reference Sequence NC_004069.1 (2002).

Durand, N. C. et al. Juicer provides a one-click system for analyzing loop-resolution Hi-C experiments. Cell Syst. 3, 95–98 (2016).

Dudchenko, O. et al. De novo assembly of the Aedes aegypti genome using Hi-C yields chromosome-length scaffolds. Science 356, 92–95 (2017).

Durand, N. C. et al. Juicebox provides a visualization system for Hi-C contact maps with unlimited zoom. Cell Syst. 3, 99–101 (2016).

Dudchenko, O. et al. The Juicebox Assembly Tools module facilitates de novo assembly of mammalian genomes with chromosome-length scaffolds for under $1000. bioRxiv https://doi.org/10.1101/254797 (2018).

Kajitani, R. et al. Efficient de novo assembly of highly heterozygous genomes from whole-genome shotgun short reads. Genome Res. 24, 1384–1395 (2014).

Shen, W., Le, S., Li, Y. & Hu, F. SeqKit: a cross-platform and ultrafast toolkit for FASTA/Q file manipulation. PLoS ONE 11, e0163962 (2016).

Murmann, A. E. et al. Comparative gene mapping in cattle, Indian muntjac, and Chinese muntjac by fluorescence in situ hybridization. Genetica 134, 345–351 (2008).

Green, R. J. & Bahr, G. F. Comparison of G-, Q-, and EM-banding patterns exhibited by the chromosome complement of the Indian muntjac, Muntiacus muntjak, with reference to nuclear DNA content and chromatin ultrastructure. Chromosoma 50, 53–67 (1975).

Carrano, A. V. et al. Purification of the chromosomes of the Indian muntjac by flow sorting. J. Histochem. Cytochem. 24, 348–354 (1976).

Session, A. M. et al. Genome evolution in the allotetraploid frog Xenopus laevis. Nature 538, 336–343 (2016).

Li, H. & Durbin, R. Fast and accurate short read alignment with Burrows-Wheeler transform. Bioinformatics 25, 1754–1760 (2009).

Smit, A. F. A. & Hubley, R. RepeatModeler Open-1.0. http://www.repeatmasker.org (2015).

Bao, W., Kojima, K. K. & Kohany, O. Repbase Update, a database of repetitive elements in eukaryotic genomes. Mob. DNA 6, 11 (2015).

Smit, A. F. A., Hubley, R. & Green, P. RepeatMasker Open-4.0. http://www.repeatmasker.org (2015).

Keilwagen, J. et al. Using intron position conservation for homology-based gene prediction. Nucleic Acids Res. 44, e89 (2016).

Cunningham, F. et al. Ensembl 2019. Nucleic Acids Res. 47, D745–D751 (2019).

Kent, W. J., Baertsch, R., Hinrichs, A., Miller, W. & Haussler, D. Evolution’s cauldron: duplication, deletion, and rearrangement in the mouse and human genomes. Proc. Natl Acad. Sci. USA 100, 11484–11489 (2003).

Li, Z. et al. Draft genomic data of the reindeer (Rangifer tarandus). GigaScience Database. https://doi.org/10.5524/100370 (2017).

Wang, Y., Coleman-Derr, D., Chen, G. & Gu, Y. Q. OrthoVenn: a web server for genome wide comparison and annotation of orthologous clusters across multiple species. Nucleic Acids Res. 43, W78–W84 (2015).

Nielsen, R. & Yang, Z. Likelihood models for detecting positively selected amino acid sites and applications to the HIV-1 envelope gene. Genetics 148, 929–936 (1998).

Larkin, M. A. et al. Clustal W and Clustal X version 2.0. Bioinformatics 23, 2947–2948 (2007).

Yang, Z. PAML 4: phylogenetic analysis by maximum likelihood. Mol. Biol. Evol. 24, 1586–1591 (2007).

The UniProt Consortium. UniProt: the universal protein knowledgebase. Nucleic Acids Res. 45, D158–D169 (2017).

Paten, B. et al. Cactus: algorithms for genome multiple sequence alignment. Genome Res. 21, 1512–1528 (2011).

Hickey, G., Paten, B., Earl, D., Zerbino, D. & Haussler, D. HAL: a hierarchical format for storing and analyzing multiple genome alignments. Bioinformatics 29, 1341–1342 (2013).

Suarez, H. G., Langer, B. E., Ladde, P. & Hiller, M. chainCleaner improves genome alignment specificity and sensitivity. Bioinformatics 33, 1596–1603 (2017).

Blanchette, M. et al. Aligning multiple genomic sequences with the threaded blockset aligner. Genome Res. 14, 708–715 (2004).

Junier, T. & Zdobnov, E. M. The Newick utilities: high-throughput phylogenetic tree processing in the UNIX shell. Bioinformatics 26, 1669–1670 (2010).

Arnold, C., Matthews, L. J. & Nunn, C. L. The 10kTrees website: a new online resource for primate phylogeny. Evol. Anthropol. 19, 114–118 (2010).

Kielbasa, S. M., Wan, R., Sato, K., Horton, P. & Frith, M. C. Adaptive seeds tame genomic sequence comparison. Genome Res. 21, 487–493 (2011).

Stamatakis, A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics 30, 1312–1313 (2014).

Kumar, S., Stecher, G. & Tamura, K. MEGA7: Molecular Evolutionary Genetics Analysis version 7.0 for bigger datasets. Mol. Biol. Evol. 33, 1870–1874 (2016).

Mello, B. Estimating timetrees with MEGA and the TimeTree resource. Mol. Biol. Evol. 35, 2334–2342 (2018).

Tamura, K. et al. Estimating divergence times in large molecular phylogenies. Proc. Natl Acad. Sci. USA 109, 19333–19338 (2012).

Krzywinski, M. et al. Circos: an information aesthetic for comparative genomics. Genome Res. 19, 1639–1645 (2009).

Li, H. et al. The Sequence Alignment/Map format and SAMtools. Bioinformatics 25, 2078–2079 (2009).


Numerical chromosomal aberrations

Each species of an organism has a specific number of chromosomes in its somatic cells. These chromosomes are found in pairs. At the time of formation of gametes the chromosome number is reduced. Hence, the gemetes carry haploid set of chromosomes. Alterations in the number of chromosomes from the diploid set is called numerical chromosomal aberration. It is also known as ploidy. There are two types of ploidy x euploidy and aneuploidy.

Euploidy is the variation in the chromosome number that occurs due to increase or decrease of full set of chromosomes. Monoploidy, diploidy and polyploidy are the types in euploidy.

In most of the plants and animals, the somatic cells contain two sets of chromosome. Diploidy is formed by the union of two gametes during fertilization.

Addition of one or more sets of chromosomes to the diploid set results in polyploidy. It is commonly noticed in plants and rare in animals. They are of two kinds - autopolyploidy and allopolyploidy.

Addition of one or more haploid set of its own genome in an organism results in autopolyploidy. Watermelon, grapes and banana are autotriploids, whereas apple is an autotetraploid.

Increase in one or more haploid set of chromosomes from two different species result in allopolyploidy. Triticale is the first man made cereal. It is obtained by crossing a wheat Triticum durum (2n = 4x = 28) and a rye

Secale cereale (2n = 2x = 14). The F 1 hybrid (2n = 3x = 21) is sterile. Then the chromosome number is doubled using colchicine and it becomes an hexaploid.

Variation that involves one or two chromosomes within the diploid set of an organism results in aneuploidy. It is of two types - hypoploidy and hyperploidy.

Decrease in one or two chromosomes from the diploid set is described as hypoploidy. There are two types of hypoploidy - monosomy and nullisomy. Monosomy is due to loss of a chromosome from the diploid set i.e. 2n - 1. Nullisomy is the condition in which a pair of homologous chromosomes is lost from the diploid set i.e. 2n - 2.

Addition of one or two chromosomes to the diploid set of chromosome results in hyperploidy. There are two types of hyperploidy - trisomy and tetrasomy. Trisomy results due to the addition of one chromosome to diploid set of chromosomes. It is represented by 2n + 1. Trisomics are observed in Datura stramonium. Tetrasomy results due to the addition of two chromosomes to diploid set of chromosome. It is represented by 2n+2.


4 Major Types of Chromosomal Aberrations (1594 Words)

The arrangement and presence of many genes on a single chromosome provides a change in genetic information not only through change in chromosome number but also by a change in chromosome structure.

Image Courtesy : neurorexia.files.wordpress.com/2013/05/figure-1-histones-1024�.jpg

The change in chromosome is due to alteration in genetic material through loss, gain or rearrangement of a particular segment. Such changes are called chromosomal aberrations. The modification brings about chromosomal mutations. Chromosomal mutations are very rare in nature but can be created artificially by ‘X’ rays, atomic radiation and chemicals, etc.

The structural changes in chromosomes are due to breaks in chromosome, or in its cell division subunit, i.e., chromatid. Each break produces 2 ends which may then follow three different paths. (Fig.43.1).

(a) They may reunite, leading to eventual loss of that chromosomal segment which does not contain the centromere.

(b) Immediate reunion or reconstitution of the same broken ends may occur, leading to reconstitution of the original structure.

(c) One or both ends of one particular break may join those produced by a different break causing an exchange, or non reconstitutional union.

Mc Clintock (1941) studied in Zea Mays that chromosome breaks and duplication follows. A dicentric chromatid is found. During anaphase spindle fibres are attached to the two centromeres resulting in the formation of bridge from one pole to other. The bridge breaks causing deficiency or duplication.

Chromosomal aberrations are of 4 major types:

(a) Deletion (b) duplication (c) inversion and (d) translocation. (Fig. 43.2).

(A) Deletion or Deficiency:

Deletion or deficiency as the name suggests there is a loss of segment of chromosome. After break the part without centromere is lost. On the other hand the part attached to the centromere acts as deficient chromosome. Bridges (1917) for the first time observed deficiency in the Bar locus of Drosophila.

Two types of deletions are found:

A single break near the end of the chromosome. Described in maize but otherwise not common.

Interstitial deletion:

Chromosome breaks and reunites but the part is lost from in between. (Fig. 43.3). Deletions are detected at the time of homologous pairing. If a part of chromosome is missing then the other chromosome also has to omit it in the form of bulging in order to make synapse. e.g., if a chromosome has 1, 2, 3, 4, genes. The part 2 is missing from one chromosome leaving, 1, 3, 4. The other homologous chromosome at the time of synapse bulge out or form loop at position 2.

If the missing segment is of physiological importance the individual will not survive. If dominant gene ‘A’ is missing the recessive allele ‘a’ may express itself. It is called pseudo dominance.

In human, deletion of chromosome 5 results in cri-du-chat syndrome, children cry like cat, they have small head and are mentally retarded.

Partial deletion of 18th chromosome results in a syndrome with large ears and long fingers.

In corn the deficiency is restricted to pollen sterility. The male haploid gemetophyte shows deficiency while female of it may receive metabolites from maternal tissue supplementing the deficiency. The omitted segment forms buckles. (Fig. 43.4)

Deficiency in E. coli is also noted. The deletion points that the DNA is single stranded and looks like collapsed loop or brush. (Fig. 43.5).

(B) Duplication:

Here a segment of chromosome is repeated twice, i.e., duplicated. Duplication was discovered in Drosophila ‘X’ chromosome for the first time carrying wild type allele for vermilion (v + ) and has been transposed to an ‘X’ chromosome carrying the mutant vermilion allele (v), Bridges found that due to the fact that ‘X’ chromosome was carrying allele v and v + both it was wild type instead of vermilion. Equal properties of v and v + produced wild type effect. Such ‘duplication females’ when crossed with nonduplicated vermilion males all female progeny was vermilion and all male progeny, i.e., y was wild type. (Fig.43.6.)

Duplication is of various types. (Fig. 43.7)

When the duplicating segment is near the centromeres e.g., the sequence on chromosome isabcdefghithe centromere is present between e and f the segment d e is repeated immediately after its normal position.

When the segment is reversed in duplication, e.g., it is d e segment that is duplicated it will be duplicated as d e e d instead of d e d e.

The segment is repeated somewhere away from its original location but on the same arm (homobrachial displacement) or on the other arm (heterobrachial displacement).

When the segment is duplicated on the non homologous chromosome it is called transposition.

Duplication involves centromere it is called extra chromosomal. In salivary gland chromosome duplications are common either as buckling in the duplication heterozygote or as cross pairing between sections of different chromosomes.

(C) Translocation:

Transfer of a section of one chromosome to non homologous chromosome is known as translocation. When there is exchange of segments on two non homologous chromosomes it is called reciprocal translocation. It also includes exchange of segments between non homologous parts of a pair of chromosomes, e.g., ‘X’ or ‘Y’ chromosomes. The segment is neither lost or added it is just exchanged.

It was first observed in Drosophila by unusual behaviour of a particular 2nd chromosome gene called Pale. It is lethal in homozygous condition. Bridges observed that its lethality can be suppressed by the presence of another gene on the 3rd chromosome which was also lethal in homozygous condition. Pale effect was caused due to deficiency for a small tip of 2nd chromosome including and plexus or balloon which links to 3rd chromosome gene between ebony or rough.

Stern in 1926 observed translocation of some allele (bobbed) on the ‘Y’ chromosome to the ‘X’ chromosome. (Fig.43.8)

Types of translocation:

(a) Simple translocation:

A single break in the chromosome and it is transferred onto the end of the other. (Fig. 43.9)

(b) Shift or intercalary translocation:

Common type of translocation involving 3 breaks so that a two break section of one chromosome (e.g., Pale) is inserted within the break produced in a non homologous chromosome. (Fig. 43.9B)

(c) Reciprocal translocation or Interchange:

Frequently observed translocation where single break in two homologous chromosomes produces an exchange of chromosome segment between them. (Fig. 43.9c)

Translocation homozygote forms the same number of homologous pairs as the normal homozygote as long as centromere is not lost.

The result of pairing and meiosis are different in translocation heterozygote bearing two translocated segments and their normal counterpart. (Fig. 43.10). A reciprocal translocation forms a 4 chromosome complex at the pachytene stage. The chiasmata between such chromosomes may form a quadrivalent which can then disjoin in 3 different segregation patterns in the first meiotic division. (Fig. 43.11).

(i) Alternate segregation:

Opposite or alternate nonhomologous centromeres go to the same pole in a zigzag fashion, so that the nontranslocated (1, 2) and translocated (1′, 2′) chromosomes are in separate gametes. Gametes have complete balanced complement of genes without duplication or deficiency (Fig. 43.11).

(ii) Adjacent-1 segregation:

Non homologous adjacent chromosomes go to the same pole but each gamete contains both translocated and non translocated chromosome (1 2′, 1𔃼) both duplication deficiencies in each gametes are present (Fig. 43.11).

(iii) Adjacent-2 segregation:

Adjacent centromeres again go to the same pole but these are now homologous as well as containing both translocated and non translocated chromosomes (1, 1′ 2, 2′). Duplication and deficiencies produce unbalanced components of gene (Fig. 43.11C).

Adjacent-1 and adjacent-2 segregation produce unbalanced gametes. Fertile gametes of translocation heterozygotes will be mostly restricted to alternate segregation.

Synapsis of heterozygous translocation chromosomes showing cross like configuration later on opens out a ring or a figure of eight (Fig. 43.12).

Consequences of such segregation are that independent assortment between genes and nonhomologous chromosomes will be inhibited. Because of duplications and deficiencies neither of the single mutant phenotypes will appear in the offspring. Translocation heterozygotes have low fertility. If extent of duplication and deficiency is small, unbalanced gametes or zygotes may not necessarily be lethal.

(D) Inversions:

A section of the chromosome becomes changed by rotation at 180° is called inversion. The order of the genes in it are reversed.

Inversion arises by the formation of loops on a chromosome. Breaks may occur at the point of intersection of the loops (Fig. 43.13). Reunion of the broken ends takes place in a new combination, and inverts. Inversion heterozygotes are formed by loops and bulges in pairs.

Paracentric inversion: The inverted segment doer no include centromere. Pericentric inversion: Inverted segments include centromere.

Paracentric inversion:

A single crossing over inverted region will result into formation of a dicentric chromosome (with 2 centromeres) and an acentric chromosome (with no centromere). Of the remaining 2 chromatids one will be normal and the other will carry inversion. The dicentric chromatid and acentric chromatid will be observed at anaphase I in the form of a bridge and a fragment (Fig. 43.14). Double crossover shows deficiencies and duplication (Fig. 43.15) giving rise to variations in anaphase I configurations.

Pericentric inversion:

In pericentric inversion centromere is in the inverted segments. In pachytene stage 2 of the 4 chromatids resulting after meiosis will have deficiencies and duplications. No dicentric bridge or acentric fragment will be observed (Fig.43.16). In pericentric inversion, if two breaks are not situated equidistant from the centromere, a change in shape of chromosome results. A metacentric chromosome may become submetacentric and vice versa (Fig. 43.17).


References

Fousteri, M. I. & Lehmann, A. R. A novel SMC protein complex in Schizosaccharomyces pombe contains the Rad18 DNA repair protein. EMBO J. 19, 1691–1702 (2000).

Guacci, V., Koshland, D. & Strunnikov, A. A direct link between sister chromatid cohesion and chromosome condensation revealed through the analysis of MCD1 in S. cerevisiae. Cell 91, 47–57 (1997).

Hirano, T., Kobayashi, R. & Hirano, M. Condensins, chromosome condensation protein complexes containing XCAP-C, XCAP-E and a Xenopus homolog of the Drosophila Barren protein. Cell 89, 511–521 (1997).

Michaelis, C., Ciosk, R. & Nasmyth, K. Cohesins: chromosomal proteins that prevent premature separation of sister chromatids. Cell 91, 35–45 (1997).

Birkenbihl, R. P. & Subramani, S. The rad21 gene product of Schizosaccharomyces pombe is a nuclear, cell cycle-regulated phosphoprotein. J. Biol. Chem. 270, 7703–7711 (1995).

Tonkin, E. T., Wang, T. J., Lisgo, S., Bamshad, M. J. & Strachan, T. NIPBL, encoding a homolog of fungal Scc2-type sister chromatid cohesion proteins and fly Nipped-B, is mutated in Cornelia de Lange syndrome. Nature Genet. 36, 636–641 (2004).

Chuang, P. T., Albertson, D. G. & Meyer, B. J. DPY-27:a chromosome condensation protein homolog that regulates C. elegans dosage compensation through association with the X chromosome. Cell 79, 459–474 (1994).

Gallego-Paez, L. M. et al. Smc5/6-mediated regulation of replication progression contributes to chromosome assembly during mitosis in human cells. Mol. Biol. Cell 25, 302–317 (2014).

Kegel, A. et al. Chromosome length influences replication-induced topological stress. Nature 471, 392–396 (2011).

Torres-Rosell, J. et al. Anaphase onset before complete DNA replication with intact checkpoint responses. Science 315, 1411–1415 (2007).

Misulovin, Z. et al. Association of cohesin and Nipped-B with transcriptionally active regions of the Drosophila melanogaster genome. Chromosoma 117, 89–102 (2008).

Kimura, K. & Hirano, T. ATP-dependent positive supercoiling of DNA by 13S condensin: a biochemical implication for chromosome condensation. Cell 90, 625–634 (1997).

Kimura, K., Rybenkov, V. V., Crisona, N. J., Hirano, T. & Cozzarelli, N. R. 13S condensin actively reconfigures DNA by introducing global positive writhe: implications for chromosome condensation. Cell 98, 239–248 (1999).

Losada, A. & Hirano, T. Intermolecular DNA interactions stimulated by the cohesin complex in vitro: implications for sister chromatid cohesion. Curr. Biol. 11, 268–272 (2001).

Hirano, T. SMC-mediated chromosome mechanics: a conserved scheme from bacteria to vertebrates? Genes Dev. 13, 11–19 (1999).

Strick, T. R., Kawaguchi, T. & Hirano, T. Real-time detection of single-molecule DNA compaction by condensin I. Curr. Biol. 14, 874–880 (2004).

Ciosk, R. et al. Cohesin's binding to chromosomes depends on a separate complex consisting of Scc2 and Scc4 proteins. Mol. Cell 5, 243–254 (2000).

Murayama, Y. & Uhlmann, F. Biochemical reconstitution of topological DNA binding by the cohesin ring. Nature 505, 367–371 (2013).

Arumugam, P. et al. ATP hydrolysis is required for cohesin's association with chromosomes. Curr. Biol. 13, 1941–1953 (2003).

Hu, B. et al. ATP hydrolysis is required for relocating cohesin from sites occupied by its Scc2/4 loading complex. Curr. Biol. 21, 12–24 (2011).

Weitzer, S., Lehane, C. & Uhlmann, F. A model for ATP hydrolysis-dependent binding of cohesin to DNA. Curr. Biol. 13, 1930–1940 (2003).

Stray, J. E., Crisona, N. J., Belotserkovskii, B. P., Lindsley, J. E. & Cozzarelli, N. R. The Saccharomyces cerevisiae Smc2/4 condensin compacts DNA into (+) chiral structures without net supercoiling. J. Biol. Chem. 280, 34723–34734 (2005).

Stray, J. E. & Lindsley, J. E. Biochemical analysis of the yeast condensin Smc2/4 complex: an ATPase that promotes knotting of circular DNA. J. Biol. Chem. 278, 26238–26248 (2003).

Sun, M., Nishino, T. & Marko, J. F. The SMC1-SMC3 cohesin heterodimer structures DNA through supercoiling-dependent loop formation. Nucleic Acids Res. 41, 6149–6160 (2013).

Anderson, D. E., Losada, A., Erickson, H. P. & Hirano, T. Condensin and cohesin display different arm conformations with characteristic hinge angles. J. Cell Biol. 156, 419–424 (2002).

Gruber, S., Haering, C. H. & Nasmyth, K. Chromosomal cohesin forms a ring. Cell 112, 765–777 (2003).

Haering, C. H., Lowe, J., Hochwagen, A. & Nasmyth, K. Molecular architecture of SMC proteins and the yeast cohesin complex. Mol. Cell 9, 773–788 (2002).

Uhlmann, F., Lottspeich, F. & Nasmyth, K. Sister-chromatid separation at anaphase onset is promoted by cleavage of the cohesin subunit Scc1. Nature 400, 37–42 (1999).

Uhlmann, F., Wernic, D., Poupart, M. A., Koonin, E. V. & Nasmyth, K. Cleavage of cohesin by the CD clan protease separin triggers anaphase in yeast. Cell 103, 375–386 (2000).

Haering, C. H., Farcas, A. M., Arumugam, P., Metson, J. & Nasmyth, K. The cohesin ring concatenates sister DNA molecules. Nature 454, 297–301 (2008).

Ivanov, D. & Nasmyth, K. A topological interaction between cohesin rings and a circular minichromosome. Cell 122, 849–860 (2005).

Ivanov, D. & Nasmyth, K. A physical assay for sister chromatid cohesion in vitro. Mol. Cell 27, 300–310 (2007).

Cuylen, S., Metz, J. & Haering, C. H. Condensin structures chromosomal DNA through topological links. Nature Struct. Mol. Biol. 18, 894–901 (2011).

Tanaka, T., Fuchs, J., Loidl, J. & Nasmyth, K. Cohesin ensures bipolar attachment of microtubules to sister centromeres and resists their precocious separation. Nature Cell Biol. 2, 492–499 (2000).

Hadjur, S. et al. Cohesins form chromosomal cis-interactions at the developmentally regulated IFNG locus. Nature 460, 410–413 (2009).

Kagey, M. H. et al. Mediator and cohesin connect gene expression and chromatin architecture. Nature 467, 430–435 (2010).

Nativio, R. et al. Cohesin is required for higher-order chromatin conformation at the imprinted IGF2-H19 locus. PLoS Genet. 5, e1000739 (2009).

Rollins, R. A., Morcillo, P. & Dorsett, D. Nipped-B, a Drosophila homologue of chromosomal adherins, participates in activation by remote enhancers in the cut and Ultrabithorax genes. Genetics 152, 577–593 (1999).

Kawauchi, S. et al. Multiple organ system defects and transcriptional dysregulation in the Nipbl( +/− ) mouse, a model of Cornelia de Lange Syndrome. PLoS Genet. 5, e1000650 (2009).

Krantz, I. D. et al. Cornelia de Lange syndrome is caused by mutations in NIPBL, the human homolog of Drosophila melanogaster Nipped-B. Nature Genet. 36, 631–635 (2004).

Parelho, V. et al. Cohesins functionally associate with CTCF on mammalian chromosome arms. Cell 132, 422–433 (2008).

Rubio, E. D. et al. CTCF physically links cohesin to chromatin. Proc. Natl Acad. Sci. USA 105, 8309–8314 (2008).

Stedman, W. et al. Cohesins localize with CTCF at the KSHV latency control region and at cellular c-myc and H19/Igf2 insulators. EMBO J. 27, 654–666 (2008).

Wendt, K. S. et al. Cohesin mediates transcriptional insulation by CCCTC-binding factor. Nature 451, 796–801 (2008).

Lopez-Serra, L., Lengronne, A., Borges, V., Kelly, G. & Uhlmann, F. Budding yeast Wapl controls sister chromatid cohesion maintenance and chromosome condensation. Curr. Biol. 23, 64–69 (2013).

Tedeschi, A. et al. Wapl is an essential regulator of chromatin structure and chromosome segregation. Nature 501, 564–568 (2013).

Gerlich, D., Koch, B., Dupeux, F., Peters, J. M. & Ellenberg, J. Live-cell imaging reveals a stable cohesin-chromatin interaction after but not before DNA replication. Curr. Biol. 16, 1571–1578 (2006).

Heidinger-Pauli, J. M., Mert, O., Davenport, C., Guacci, V. & Koshland, D. Systematic reduction of cohesin differentially affects chromosome segregation, condensation, and DNA repair. Curr. Biol. 20, 957–963 (2010).

Kaur, M. et al. Precocious sister chromatid separation (PSCS) in Cornelia de Lange syndrome. Am. J. Med. Genet. A 138A, 27–31 (2005).

Remeseiro, S. et al. Reduction of Nipbl impairs cohesin loading locally and affects transcription but not cohesion-dependent functions in a mouse model of Cornelia de Lange Syndrome. Biochim. Biophys. Acta 1832, 2097–2102 (2013).

Cui, Y., Petrushenko, Z. M. & Rybenkov, V. V. MukB acts as a macromolecular clamp in DNA condensation. Nature Struct. Mol. Biol. 15, 411–418 (2008).

Hirota, T., Gerlich, D., Koch, B., Ellenberg, J. & Peters, J. M. Distinct functions of condensin I and II in mitotic chromosome assembly. J. Cell Sci. 117, 6435–6445 (2004).

Ono, T., Fang, Y., Spector, D. L. & Hirano, T. Spatial and temporal regulation of Condensins I and II in mitotic chromosome assembly in human cells. Mol. Biol. Cell 15, 3296–3308 (2004).

Maeshima, K. & Laemmli, U. K. A two-step scaffolding model for mitotic chromosome assembly. Dev. Cell 4, 467–480 (2003).

Wang, J. C. Cellular roles of DNA topoisomerases: a molecular perspective. Nature Rev. Mol. Cell Biol. 3, 430–440 (2002).

Bhalla, N., Biggins, S. & Murray, A. W. Mutation of YCS4, a budding yeast condensin subunit, affects mitotic and nonmitotic chromosome behavior. Mol. Biol. Cell 13, 632–645 (2002).

Bhat, M. A., Philp, A. V., Glover, D. M. & Bellen, H. J. Chromatid segregation at anaphase requires the barren product, a novel chromosome-associated protein that interacts with Topoisomerase II. Cell 87, 1103–1114 (1996).

Coelho, P. A., Queiroz-Machado, J. & Sunkel, C. E. Condensin-dependent localisation of topoisomerase II to an axial chromosomal structure is required for sister chromatid resolution during mitosis. J. Cell Sci. 116, 4763–4776 (2003).

Hudson, D. F., Vagnarelli, P., Gassmann, R. & Earnshaw, W. C. Condensin is required for nonhistone protein assembly and structural integrity of vertebrate mitotic chromosomes. Dev. Cell 5, 323–336 (2003).

Baxter, J. et al. Positive supercoiling of mitotic DNA drives decatenation by topoisomerase II in eukaryotes. Science 331, 1328–1332 (2011).

Charbin, A., Bouchoux, C. & Uhlmann, F. Condensin aids sister chromatid decatenation by topoisomerase II. Nucleic Acids Res. 42, 340–348 (2014).

D'Ambrosio, C., Kelly, G., Shirahige, K. & Uhlmann, F. Condensin-dependent rDNA decatenation introduces a temporal pattern to chromosome segregation. Curr. Biol. 18, 1084–1089 (2008).

Lieb, J. D., Albrecht, M. R., Chuang, P. T. & Meyer, B. J. MIX-1: an essential component of the C. elegans mitotic machinery executes X chromosome dosage compensation. Cell 92, 265–277 (1998).

Meyer, B. J. Targeting X chromosomes for repression. Curr. Opin. Genet. Dev. 20, 179–189 (2010).

Rawlings, J. S., Gatzka, M., Thomas, P. G. & Ihle, J. N. Chromatin condensation via the condensin II complex is required for peripheral T-cell quiescence. EMBO J. 30, 263–276 (2011).

Duncan, I. W. Transvection effects in Drosophila. Annu. Rev. Genet. 36, 521–556 (2002).

Hartl, T. A., Smith, H. F. & Bosco, G. Chromosome alignment and transvection are antagonized by condensin II. Science 322, 1384–1387 (2008).

Almedawar, S., Colomina, N., Bermudez-Lopez, M., Pocino-Merino, I. & Torres-Rosell, J. A. SUMO-dependent step during establishment of sister chromatid cohesion. Curr. Biol. 22, 1576–1581 (2012).

Stephan, A. K., Kliszczak, M., Dodson, H., Cooley, C. & Morrison, C. G. Roles of vertebrate Smc5 in sister chromatid cohesion and homologous recombinational repair. Mol. Cell. Biol. 31, 1369–1381 (2011).

McAleenan, A. et al. SUMOylation of the alpha- kleisin subunit of cohesin is required for DNA damage-induced cohesion. Curr. Biol. 22, 1564–1575 (2012).

Potts, P. R. & Yu, H. The SMC5/6 complex maintains telomere length in ALT cancer cells through SUMOylation of telomere-binding proteins. Nature Struct. Mol. Biol. 14, 581–590 (2007).

Takahashi, Y. et al. Cooperation of sumoylated chromosomal proteins in rDNA maintenance. PLoS Genet. 4, e1000215 (2008).

Wu, N. et al. Scc1 sumoylation by Mms21 promotes sister chromatid recombination through counteracting Wapl. Genes Dev. 26, 1473–1485 (2012).

Zhao, X. & Blobel, G. A. SUMO ligase is part of a nuclear multiprotein complex that affects DNA repair and chromosomal organization. Proc. Natl Acad. Sci. USA 102, 4777–4782 (2005).

Zhao, X. & Blobel, G. From The Cover: A SUMO ligase is part of a nuclear multiprotein complex that affects DNA repair and chromosomal organization. Proc. Natl Acad. Sci. USA 102, 4777–4782 (2005).

Wolters, S. et al. Loss of Caenorhabditis elegans BRCA1 promotes genome stability during replication in smc-5 mutants. Genetics 196, 985–999 (2014).

Lindroos, H. B. et al. Chromosomal association of the Smc5/6 complex reveals that it functions in differently regulated pathways. Mol. Cell 22, 755–767 (2006).

Pflumm, M. F. & Botchan, M. R. Orc mutants arrest in metaphase with abnormally condensed chromosomes. Development 128, 1697–1707 (2001).

Bavner, A., Matthews, J., Sanyal, S., Gustafsson, J. A. & Treuter, E. EID3 is a novel EID family member and an inhibitor of CBP-dependent co-activation. Nucleic Acids Res. 33, 3561–3569 (2005).

Taylor, E. M., Copsey, A. C., Hudson, J. J., Vidot, S. & Lehmann, A. R. Identification of the proteins, including MAGEG1, that make up the human SMC5-6 protein complex. Mol. Cell. Biol. 28, 1197–1206 (2008).

Nasmyth, K. Cohesin: a catenase with separate entry and exit gates? Nature Cell Biol. 13, 1170–1177 (2011).

Gillespie, P. J. & Hirano, T. Scc2 couples replication licensing to sister chromatid cohesion in Xenopus egg extracts. Curr. Biol. 14, 1598–1603 (2004).

Takahashi, T. S., Yiu, P., Chou, M. F., Gygi, S. & Walter, J. C. Recruitment of Xenopus Scc2 and cohesin to chromatin requires the pre-replication complex. Nature Cell Biol. 6, 991–996 (2004).

Watrin, E. et al. Human Scc4 is required for cohesin binding to chromatin, sister-chromatid cohesion, and mitotic progression. Curr. Biol. 16, 863–874 (2006).

Gandhi, R., Gillespie, P. J. & Hirano, T. Human Wapl is a cohesin-binding protein that promotes sister-chromatid resolution in mitotic prophase. Curr. Biol. 16, 2406–2417 (2006).

Kueng, S. et al. Wapl controls the dynamic association of cohesin with chromatin. Cell 127, 955–967 (2006).

Rolef Ben-Shahar, T. et al. Eco1-dependent cohesin acetylation during establishment of sister chromatid cohesion. Science 321, 563–566 (2008).

Rowland, B. D. et al. Building sister chromatid cohesion: smc3 acetylation counteracts an antiestablishment activity. Mol. Cell 33, 763–774 (2009).

Sutani, T., Kawaguchi, T., Kanno, R., Itoh, T. & Shirahige, K. Budding yeast Wpl1(Rad61)-Pds5 complex counteracts sister chromatid cohesion-establishing reaction. Curr. Biol. 19, 492–497 (2009).

Unal, E. et al. A molecular determinant for the establishment of sister chromatid cohesion. Science 321, 566–569 (2008).

Zhang, J. et al. Acetylation of Smc3 by Eco1 is required for S phase sister chromatid cohesion in both human and yeast. Mol. Cell 31, 143–151 (2008).

Lafont, A. L., Song, J. & Rankin, S. Sororin cooperates with the acetyltransferase Eco2 to ensure DNA replication-dependent sister chromatid cohesion. Proc. Natl Acad. Sci. USA 107, 20364–20369 (2010).

Nishiyama, T. et al. Sororin mediates sister chromatid cohesion by antagonizing Wapl. Cell 143, 737–749 (2010).

Rankin, S., Ayad, N. G. & Kirschner, M. W. Sororin, a substrate of the anaphase-promoting complex, is required for sister chromatid cohesion in vertebrates. Mol. Cell 18, 185–200 (2005).

Uhlmann, F. & Nasmyth, K. Cohesion between sister chromatids must be established during DNA replication. Curr. Biol. 8, 1095–1101 (1998).

Lengronne, A. et al. Establishment of sister chromatid cohesion at the S. cerevisiae replication fork. Mol. Cell 23, 787–799 (2006).

Mayer, M. L., Gygi, S. P., Aebersold, R. & Hieter, P. Identification of RFC(Ctf18p, Ctf8p, Dcc1p): an alternative RFC complex required for sister chromatid cohesion in S. cerevisiae. Mol. Cell 7, 959–970 (2001).

Moldovan, G. L., Pfander, B. & Jentsch, S. PCNA controls establishment of sister chromatid cohesion during S phase. Mol. Cell 23, 723–732 (2006).

Skibbens, R. V. Chl1p, a DNA helicase-like protein in budding yeast, functions in sister-chromatid cohesion. Genetics 166, 33–42 (2004).

Song, J. et al. Cohesin acetylation promotes sister chromatid cohesion only in association with the replication machinery. J. Biol. Chem. 287, 34325–34336 (2012).

Strom, L. et al. Postreplicative formation of cohesion is required for repair and induced by a single DNA break. Science 317, 242–245 (2007).

Unal, E., Heidinger-Pauli, J. M. & Koshland, D. DNA double-strand breaks trigger genome-wide sister-chromatid cohesion through Eco1 (Ctf7). Science 317, 245–248 (2007).

Lyons, N. A. & Morgan, D. O. Cdk1-dependent destruction of Eco1 prevents cohesion establishment after S phase. Mol. Cell 42, 378–389 (2011).

Hauf, S., Waizenegger, I. C. & Peters, J. M. Cohesin cleavage by separase required for anaphase and cytokinesis in human cells. Science 293, 1320–1323 (2001).

Schmidt, D. et al. A CTCF-independent role for cohesin in tissue-specific transcription. Genome Res. 20, 578–588 (2010).

Liu, J. et al. Transcriptional dysregulation in NIPBL and cohesin mutant human cells. PLoS Biol. 7, e1000119 (2009).

Pauli, A. et al. A direct role for cohesin in gene regulation and ecdysone response in Drosophila salivary glands. Curr. Biol. 20, 1787–1798 (2010).

Schaaf, C. A. et al. Regulation of the Drosophila Enhancer of split and invected-engrailed gene complexes by sister chromatid cohesion proteins. PLoS ONE 4, e6202 (2009).

Zuin, J. et al. Cohesin and CTCF differentially affect chromatin architecture and gene expression in human cells. Proc. Natl Acad. Sci. USA 111, 996–1001 (2014).

Laloraya, S., Guacci, V. & Koshland, D. Chromosomal addresses of the cohesin component Mcd1p. J. Cell Biol. 151, 1047–1056 (2000).

Lengronne, A. et al. Cohesin relocation from sites of chromosomal loading to places of convergent transcription. Nature 430, 573–578 (2004).

Tanaka, T., Cosma, M. P., Wirth, K. & Nasmyth, K. Identification of cohesin association sites at centromeres and along chromosome arms. Cell 98, 847–858 (1999).

Bernard, P. et al. Requirement of heterochromatin for cohesion at centromeres. Science 294, 2539–2542 (2001).

Gullerova, M. & Proudfoot, N. J. Cohesin complex promotes transcriptional termination between convergent genes in S. pombe. Cell 132, 983–995 (2008).

Schmidt, C. K., Brookes, N. & Uhlmann, F. Conserved features of cohesin binding along fission yeast chromosomes. Genome Biol. 10, R52 (2009).

Deardorff, M. A. et al. HDAC8 mutations in Cornelia de Lange syndrome affect the cohesin acetylation cycle. Nature 489, 313–317 (2012).

Ono, T. et al. Differential contributions of condensin I and condensin II to mitotic chromosome architecture in vertebrate cells. Cell 115, 109–121 (2003).

Gerlich, D., Hirota, T., Koch, B., Peters, J. M. & Ellenberg, J. Condensin I stabilizes chromosomes mechanically through a dynamic interaction in live cells. Curr. Biol. 16, 333–344 (2006).

Oliveira, R. A., Heidmann, S. & Sunkel, C. E. Condensin I binds chromatin early in prophase and displays a highly dynamic association with Drosophila mitotic chromosomes. Chromosoma 116, 259–274 (2007).

Hirano, T. Condensins: universal organizers of chromosomes with diverse functions. Genes Dev. 26, 1659–1678 (2012).

Shintomi, K. & Hirano, T. The relative ratio of condensin I to II determines chromosome shapes. Genes Dev. 25, 1464–1469 (2011).

Lipp, J. J., Hirota, T., Poser, I. & Peters, J. M. Aurora B controls the association of condensin I but not condensin II with mitotic chromosomes. J. Cell Sci. 120, 1245–1255 (2007).

Takemoto, A. et al. Analysis of the role of Aurora B on the chromosomal targeting of condensin I. Nucleic Acids Res. 35, 2403–2412 (2007).

Abe, S. et al. The initial phase of chromosome condensation requires Cdk1-mediated phosphorylation of the CAP-D3 subunit of condensin II. Genes Dev. 25, 863–874 (2011).

St-Pierre, J. et al. Polo kinase regulates mitotic chromosome condensation by hyperactivation of condensin DNA supercoiling activity. Mol. Cell 34, 416–426 (2009).

D'Ambrosio, C. et al. Identification of cis-acting sites for condensin loading onto budding yeast chromosomes. Genes Dev. 22, 2215–2227 (2008).

Haeusler, R. A., Pratt-Hyatt, M., Good, P. D., Gipson, T. A. & Engelke, D. R. Clustering of yeast tRNA genes is mediated by specific association of condensin with tRNA gene transcription complexes. Genes Dev. 22, 2204–2214 (2008).

Tada, K., Susumu, H., Sakuno, T. & Watanabe, Y. Condensin association with histone H2A shapes mitotic chromosomes. Nature 474, 477–483 (2011).

Iwasaki, O., Tanaka, A., Tanizawa, H., Grewal, S. I. & Noma, K. Centromeric localization of dispersed Pol III genes in fission yeast. Mol. Biol. Cell 21, 254–265 (2010).

Kranz, A. L. et al. Genome-wide analysis of condensin binding in Caenorhabditis elegans. Genome Biol. 14, R112 (2013).

Dawes, H. E. et al. Dosage compensation proteins targeted to X chromosomes by a determinant of hermaphrodite fate. Science 284, 1800–1804 (1999).

Davis, T. L. & Meyer, B. J. SDC-3 coordinates the assembly of a dosage compensation complex on the nematode X chromosome. Development 124, 1019–1031 (1997).

Hsu, D. R. & Meyer, B. J. The dpy-30 gene encodes an essential component of the Caenorhabditis elegans dosage compensation machinery. Genetics 137, 999–1018 (1994).

Pferdehirt, R. R., Kruesi, W. S. & Meyer, B. J. An MLL/COMPASS subunit functions in the C. elegans dosage compensation complex to target X chromosomes for transcriptional regulation of gene expression. Genes Dev. 25, 499–515 (2011).

Csankovszki, G., McDonel, P. & Meyer, B. J. Recruitment and spreading of the C. elegans dosage compensation complex along X chromosomes. Science 303, 1182–1185 (2004).

Jans, J. et al. A condensin-like dosage compensation complex acts at a distance to control expression throughout the genome. Genes Dev. 23, 602–618 (2009).

McDonel, P., Jans, J., Peterson, B. K. & Meyer, B. J. Clustered DNA motifs mark X chromosomes for repression by a dosage compensation complex. Nature 444, 614–618 (2006).

Torres-Rosell, J. et al. SMC5 and SMC6 genes are required for the segregation of repetitive chromosome regions. Nature Cell Biol. 7, 412–419 (2005).

Pebernard, S., Schaffer, L., Campbell, D., Head, S. R. & Boddy, M. N. Localization of Smc5/6 to centromeres and telomeres requires heterochromatin and SUMO, respectively. EMBO J. 27, 3011–3023 (2008).

Katou, Y., Kaneshiro, K., Aburatani, H. & Shirahige, K. Genomic approach for the understanding of dynamic aspect of chromosome behavior. Methods Enzymol. 409, 389–410 (2006).

Nakato, R., Itoh, T. & Shirahige, K. DROMPA: easy-to-handle peak calling and visualization software for the computational analysis and validation of ChIP-seq data. Genes Cells 18, 589–601 (2013).

Poorey, K. et al. Measuring chromatin interaction dynamics on the second time scale at single-copy genes. Science 342, 369–372 (2013).

Zuin, J. et al. A cohesin-independent role for NIPBL at promoters provides insights in CdLS. PLoS Genet. 10, e1004153 (2014).

Auerbach, R. K. et al. Mapping accessible chromatin regions using Sono-Seq. Proc. Natl Acad. Sci. USA 106, 14926–14931 (2009).

Teytelman, L., Thurtle, D. M., Rine, J. & van Oudenaarden, A. Highly expressed loci are vulnerable to misleading ChIP localization of multiple unrelated proteins. Proc. Natl Acad. Sci. USA 110, 18602–18607 (2013).

Dekker, J., Marti-Renom, M. A. & Mirny, L. A. Exploring the three-dimensional organization of genomes: interpreting chromatin interaction data. Nature Rev. Genet. 14, 390–403 (2013).

Sergeant, J. et al. Composition and architecture of the Schizosaccharomyces pombe Rad18 (Smc5-6) complex. Mol. Cell. Biol. 25, 172–184 (2005).

Hopfner, K. P. et al. Structural biology of Rad50 ATPase: ATP-driven conformational control in DNA double-strand break repair and the ABC-ATPase superfamily. Cell 101, 789–800 (2000).

Lowe, J., Cordell, S. C. & van den Ent, F. Crystal structure of the SMC head domain: an ABC ATPase with 900 residues antiparallel coiled-coil inserted. J. Mol. Biol. 306, 25–35 (2001).

Hirano, M. & Hirano, T. Hinge-mediated dimerization of SMC protein is essential for its dynamic interaction with DNA. EMBO J. 21, 5733–5744 (2002).

Yamazoe, M. et al. Complex formation of MukB, MukE and MukF proteins involved in chromosome partitioning in Escherichia coli. EMBO J. 18, 5873–5884 (1999).

Schleiffer, A. et al. Kleisins: a superfamily of bacterial and eukaryotic SMC protein partners. Mol. Cell 11, 571–575 (2003).

Toth, A. et al. Yeast cohesin complex requires a conserved protein, Eco1p(Ctf7), to establish cohesion between sister chromatids during DNA replication. Genes Dev. 13, 320–333 (1999).

Hartman, T., Stead, K., Koshland, D. & Guacci, V. Pds5p is an essential chromosomal protein required for both sister chromatid cohesion and condensation in Saccharomyces cerevisiae. J. Cell Biol. 151, 613–626 (2000).

Panizza, S., Tanaka, T., Hochwagen, A., Eisenhaber, F. & Nasmyth, K. Pds5 cooperates with cohesin in maintaining sister chromatid cohesion. Curr. Biol. 10, 1557–1564 (2000).

Shintomi, K. & Hirano, T. Releasing cohesin from chromosome arms in early mitosis: opposing actions of Wapl-Pds5 and Sgo1. Genes Dev. 23, 2224–2236 (2009).

Chan, K. L. et al. Cohesin's DNA exit gate is distinct from its entrance gate and is regulated by acetylation. Cell 150, 961–974 (2012).

Sutani, T. et al. Fission yeast condensin complex: essential roles of non-SMC subunits for condensation and Cdc2 phosphorylation of Cut3/SMC4. Genes Dev. 13, 2271–2283 (1999).

Freeman, L., Aragon-Alcaide, L. & Strunnikov, A. The condensin complex governs chromosome condensation and mitotic transmission of rDNA. J. Cell Biol. 149, 811–824 (2000).

Amberg, D. C., Burke, D. J. & Strathern, J. N. Methods in Yeast Genetics. Cold Spring Harbor Laboratory Course Manual (Cold Spring Harbor Laboratory Press, 2005).

Nairz, K. & Klein, F. mre11S—a yeast mutation that blocks double-strand-break processing and permits nonhomologous synapsis in meiosis. Genes Dev. 11, 2272–2290 (1997).

Csankovszki, G. et al. Three distinct condensin complexes control C. elegans chromosome dynamics. Curr. Biol. 19, 9–19 (2009).

Pebernard, S. et al. The Nse5-Nse6 dimer mediates DNA repair roles of the Smc5-Smc6 complex. Mol. Cell. Biol. 26, 1617–1630 (2006).

Palecek, J., Vidot, S., Feng, M., Doherty, A. J. & Lehmann, A. R. The Smc5-Smc6 DNA repair complex. bridging of the Smc5-Smc6 heads by the KLEISIN, Nse4, and non-Kleisin subunits. J. Biol. Chem. 281, 36952–36959 (2006).

Potts, P. R. & Yu, H. Human MMS21/NSE2 is a SUMO ligase required for DNA repair. Mol. Cell. Biol. 25, 7021–7032 (2005).

Andrews, E. A. et al. Nse2, a component of the Smc5-6 complex, is a SUMO ligase required for the response to DNA damage. Mol. Cell. Biol. 25, 185–196 (2005).

Duan, X. et al. Architecture of the Smc5/6 Complex of Saccharomyces cerevisiae Reveals a Unique Interaction between the Nse5-6 Subcomplex and the Hinge Regions of Smc5 and Smc6. J. Biol. Chem. 284, 8507–8515 (2009).


Watch the video: Πως θα ξεχωρίσεις: Χρωμοσώματα - DNA - Καρυότυπο (January 2022).