Information

Are any organisms known to use meiosis I to create non-identical offspring asexually?

Are any organisms known to use meiosis I to create non-identical offspring asexually?



We are searching data for your request:

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

So, there are numerous species of animals who use parthenogenesis, but to my knowledge the reproduction is clonal. That is, the offspring are identical to the mother. Are there any documented cases where a female goes through meiosis one to produce varied cells that are now diploid and these cells do not go through meiosis two, but rather develop in to a diploid organism? It would seem an effective reproductive strategy in that it generates genetic variation without a mate (at least more-so than strict asexual reproduction). Is my logic flawed in some way?


In automixy the meiotic cells give rise to diploid offsprings. This can happen by diploidization of the haploid cell (1n->2n), which will produce homozygotes or endomitosis prior to meiosis (4n->2n) which produces heterozygotes. Examples:

  1. Cnemidophorus uniparens :4n->2n

  2. Sphyrna tiburo:1n->2n

I don't know of any case where there is fusion of similar gametes to form a diploid cell. It is difficult for two ova to fuse in natural conditions because the vitelline membrane has to be dissolved. Experimentally a haploid ES-cell can be fused to ovum to form a progeny. Haploid ES cells undergo diploidization and when injected in blastocyst, can develop properly (Ref). In fact haploid androgenic-ES cell line had been made in this study by injecting sperm into an enucleated oocyte. These androgenic-haploid cell lines can be fused to ovum to give rise to viable offsprings.

Also see this question. Similar topic


Chapter 15. Meiosis & Sexual Reproduction

Figure 15.1 Each of us, like these other large multicellular organisms, begins life as a fertilized egg. After trillions of cell divisions, each of us develops into a complex, multicellular organism. (Credit a: modification of work by Frank Wouters credit b: modification of work by Ken Cole, USGS credit c: modification of work by Martin Pettitt)
  • 15.1 The Process of Meiosis
  • 15.2 Disorders in Chromosome Number
  • 15.3 Sexual Reproduction

11.1 The Process of Meiosis

In this section, you will explore the following questions:

  • How do chromosomes behave during meiosis?
  • What cellular events occur during meiosis?
  • What are the similarities and differences between meiosis and mitosis?
  • How can the process of meiosis generate genetic variation?

Connection for AP ® Courses

As we explored the cell cycle and mitosis in a previous chapter, we learned that cells divide to grow, replace other cells, and reproduce asexually. Without mutation, or changes in the DNA, the daughter cells produced by mitosis receive a set of genetic instructions that is identical to that of the parent cell. Because changes in genes drive both the unity and diversity of life, organisms without genetic variation cannot evolve through natural selection. Evolution occurs only because organisms have developed ways to vary their genetic material. This occurs through mutations in DNA, recombination of genes during meiosis, and meiosis followed by fertilization in sexually reproducing organisms.

Sexual reproduction requires that diploid (2n) organisms produce haploid (1n) cells through meiosis and that these haploid cells fuse to form new, diploid offspring. The union of these two haploid cells, one from each parent, is fertilization. Although the processes of meiosis and mitosis share similarities, their end products are different. Recall that eukaryotic DNA is contained in chromosomes, and that chromosomes occur in homologous pairs (homologues). At fertilization, the male parent contributes one member of each homologous pair to the offspring, and the female parent contributes the other. With the exception of the sex chromosomes, homologous chromosomes contain the same genes, but these genes can have different variations, called alleles. (For example, you might have inherited an allele for brown eyes from your father and an allele for blue eyes from your mother.) As in mitosis, homologous chromosomes are duplicated during the S-stage (synthesis) of interphase. However, unlike mitosis, in which there is just one nuclear division, meiosis has two complete rounds of nuclear division—meiosis I and meiosis II. These result in four nuclei and (usually) four daughter cells, each with half the number of chromosomes as the parent cell (1n). The first division, meiosis I, separates homologous chromosomes, and the second division, meiosis II, separates chromatids. (Remember: during meiosis, DNA replicates ONCE but divides TWICE, whereas in mitosis, DNA replicates ONCE but divides only ONCE.).

Although mitosis and meiosis are similar in many ways, they have different outcomes. The main difference is in the type of cell produced: mitosis produces identical cells, allowing growth or repair of tissues meiosis generates reproductive cells, or gametes. Gametes, often called sex cells, unite with other sex cells to produce new, unique organisms.

Genetic variation occurs during meiosis I, in which homologous chromosomes pair and exchange non-sister chromatid segments (crossover). Here the homologous chromosomes separate into different nuclei, causing a reduction in “ploidy.” During meiosis II—which is more similar to a mitotic division—the chromatids separate and segregate into four haploid sex cells. However, because of crossover, the resultant daughter cells do not contain identical genomes. As in mitosis, external factors and internal signals regulate the meiotic cell cycle. As we will explore in more detail in a later chapter, errors in meiosis can cause genetic disorders, such as Down syndrome.

Information presented and the examples highlighted in the section support concepts and learning objectives outlined in Big Idea 3 of the AP ® Biology Curriculum Framework. The learning objectives listed in the Curriculum Framework provide a transparent foundation for the AP ® Biology course, an inquiry-based laboratory experience, instructional activities, and AP ® exam questions. A learning objective merges required content with one or more of the seven science practices.

Big Idea 3 Living systems store, retrieve, transmit and respond to information essential to life processes.
Enduring Understanding 3.A Heritable information provides for continuity of life.
Essential Knowledge 3.A.2 In eukaryotes, heritable information is passed to the next generation via processes that include the cell cycle and mitosis or meiosis plus fertilization.
Science Practice 6.2 The student can construct explanations of phenomena based on evidence produced through scientific practices.
Learning Objective 3.9 The student is able to construct an explanation, using visual representations or narratives, as to how DNA in chromosomes is transmitted to the next generation via mitosis, or meiosis followed by fertilization.
Essential Knowledge 3.A.2 In eukaryotes, heritable information is passed to the next generation via processes that include the cell cycle and mitosis or meiosis plus fertilization.
Science Practice 7.1 The student can connect phenomena and models across spatial and temporal scales.
Learning Objective 3.10 The student is able to represent the connection between meiosis and increased genetic diversity necessary for evolution.

The Science Practice Challenge Questions contain additional test questions for this section that will help you prepare for the AP exam. These questions address the following standards:
[APLO 1.9][APLO 2.15][APLO 2.39][APLO 3.11][APLO 3.9]

Teacher Support

The process of meiosis can be confusing, especially if it is taught as just a series of steps. Initially, discuss the goal of the process. Explain that meiosis serves to produce reproductive cells with exactly half the number of chromosomes, and that once these haploid cells are fused during fertilization, a complete set of genetic instructions for a new individual is formed. Meiosis starts in a cell with chromosomes in pairs. Each chromosome has already been duplicated and the two sister strands are held together. Therefore, each pair consists of four chromatids. Because students have already learned about mitosis (the process whereby chromosomes are sorted and allocated to daughter cells), it might be helpful to teach meiosis as a special case of mitosis. The first division separates the pairs of chromosomes, reducing the number of duplicated chromosomes in the daughter cells by half. The second division separates the chromatids, creating daughter cells that each has one half of the total number of chromosomes of the original cell. An added benefit to an organism using meiosis is the increase in genetic variation that occurs during the process. Each individual born as a result of sexual reproduction truly has a unique assortment of genes.

You read that fertilization is the union of two sex cells from two individual organisms. If these two cells each contain one set of chromosomes, the resulting fertilized cell contains two sets of chromosomes. Haploid cells contain one set of chromosomes. Cells containing two sets of chromosomes are called diploid. The number of sets of chromosomes in a cell is called its ploidy level. If the reproductive cycle is to continue, a diploid cell must reduce the number of its chromosome sets before fertilization can occur again. Otherwise, the number of chromosome sets would double, and continue to double in every generation. So, in addition to fertilization, sexual reproduction includes a nuclear division that reduces the number of chromosome sets.

Most animals and plants are diploid, containing two sets of chromosomes. In an organism’s somatic cells , sometimes referred to as “body” cells (all cells of a multicellular organism except the reproductive cells), the nucleus contains two copies of each chromosome, called homologous chromosomes. Homologous chromosomes are matched pairs containing the same genes in identical locations along their length. Diploid organisms inherit one copy of each homologous chromosome from each parent all together, they are considered a full set of chromosomes. Haploid cells, containing a single copy of each homologous chromosome, are found only within an organism’’s reproductive structures, such as the ovaries and testes. Haploid cells can be either gametes or spores. Male gametes are sperm and female gametes are eggs. All animals and most plants produce gametes. Spores are haploid cells that can produce a haploid organism or can fuse with another spore to form a diploid cell. Some plants and all fungi produce spores.

As you have learned, the nuclear division that forms haploid cells— meiosis —is closely related to mitosis. Mitosis is the part of a cell reproduction cycle that results in identical daughter nuclei that are also genetically identical to the original parent nucleus. In mitosis, both the parent and the daughter nuclei are at the same ploidy level—diploid for most plants and animals. Meiosis employs many of the same mechanisms as mitosis. However, the starting nucleus is always diploid and the nuclei that result at the end of a meiotic cell division are haploid. To achieve this reduction in chromosome number, meiosis consists of one round of chromosome duplication and two rounds of nuclear division. Because the events that occur during each of the division stages are analogous to the events of mitosis, the same stage names are assigned. However, because there are two rounds of division, the major process and the stages are designated with a “I” or a “II.” Thus, meiosis I is the first round of meiotic division and consists of prophase I, prometaphase I, and so on. Meiosis II , in which the second round of meiotic division takes place, includes prophase II, prometaphase II, and so on.

Teacher Support

Meiosis I has the same steps as mitosis, with the exception that the chromosome pairs, not the chromatids, are separated at anaphase I. Two other events occur during the first cell division to produce the genetic variation that results. In prophase I, when the pairs of chromosomes condense and tentatively join, parts of the arms and legs of the chromosomes can crossover, or exchange places, with corresponding parts on the other homologous chromosome. The resulting pair now has a configuration that was not present initially. The pairs line up in a double line during metaphase I, but the distribution of the pairs at the equator is random. Half of the original chromosomes came from one parent, half from the other. As the chromosomes line up and are pulled apart during anaphase I, each daughter cell will receive a chromosome mixture that was not present in the original germ cells. Figure 11.3 illustrates crossing over and Figure 11.4 illustrates the random distribution of pairs of chromosomes. Also use the Link to Learning: Meiosis: An Interactive Animation. Meiosis II finishes the process and closely resembles mitosis, except for the number of chromosomes present, as compared to somatic cells.

Teacher Support

Comparing meiosis and mitosis should be a review of the two processes, with a reinforcement of the similarities and differences.

Meiosis I

Meiosis is preceded by an interphase consisting of the G1, S, and G2 phases, which are nearly identical to the phases preceding mitosis. The G1 phase, which is also called the first gap phase, is the first phase of the interphase and is focused on cell growth. The S phase is the second phase of interphase, during which the DNA of the chromosomes is replicated. Finally, the G2 phase, also called the second gap phase, is the third and final phase of interphase in this phase, the cell undergoes the final preparations for meiosis.

During DNA duplication in the S phase, each chromosome is replicated to produce two identical copies, called sister chromatids, that are held together at the centromere by cohesin proteins. Cohesin holds the chromatids together until anaphase II. The centrosomes, which are the structures that organize the microtubules of the meiotic spindle, also replicate. This prepares the cell to enter prophase I, the first meiotic phase.

Prophase I

Early in prophase I, before the chromosomes can be seen clearly microscopically, the homologous chromosomes are attached at their tips to the nuclear envelope by proteins. As the nuclear envelope begins to break down, the proteins associated with homologous chromosomes bring the pair close to each other. Recall that, in mitosis, homologous chromosomes do not pair together. In mitosis, homologous chromosomes line up end-to-end so that when they divide, each daughter cell receives a sister chromatid from both members of the homologous pair. The synaptonemal complex , a lattice of proteins between the homologous chromosomes, first forms at specific locations and then spreads to cover the entire length of the chromosomes. The tight pairing of the homologous chromosomes is called synapsis . In synapsis, the genes on the chromatids of the homologous chromosomes are aligned precisely with each other. The synaptonemal complex supports the exchange of chromosomal segments between non-sister homologous chromatids, a process called crossing over. Crossing over can be observed visually after the exchange as chiasmata (singular = chiasma) (Figure 11.2).

In species such as humans, even though the X and Y sex chromosomes are not homologous (most of their genes differ), they have a small region of homology that allows the X and Y chromosomes to pair up during prophase I. A partial synaptonemal complex develops only between the regions of homology.

Located at intervals along the synaptonemal complex are large protein assemblies called recombination nodules . These assemblies mark the points of later chiasmata and mediate the multistep process of crossover —or genetic recombination—between the non-sister chromatids. Near the recombination nodule on each chromatid, the double-stranded DNA is cleaved, the cut ends are modified, and a new connection is made between the non-sister chromatids. As prophase I progresses, the synaptonemal complex begins to break down and the chromosomes begin to condense. When the synaptonemal complex is gone, the homologous chromosomes remain attached to each other at the centromere and at chiasmata. The chiasmata remain until anaphase I. The number of chiasmata varies according to the species and the length of the chromosome. There must be at least one chiasma per chromosome for proper separation of homologous chromosomes during meiosis I, but there may be as many as 25. Following crossover, the synaptonemal complex breaks down and the cohesin connection between homologous pairs is also removed. At the end of prophase I, the pairs are held together only at the chiasmata (Figure 11.3) and are called tetrads because the four sister chromatids of each pair of homologous chromosomes are now visible.

The crossover events are the first source of genetic variation in the nuclei produced by meiosis. A single crossover event between homologous non-sister chromatids leads to a reciprocal exchange of equivalent DNA between a maternal chromosome and a paternal chromosome. Now, when that sister chromatid is moved into a gamete cell it will carry some DNA from one parent of the individual and some DNA from the other parent. The sister recombinant chromatid has a combination of maternal and paternal genes that did not exist before the crossover. Multiple crossovers in an arm of the chromosome have the same effect, exchanging segments of DNA to create recombinant chromosomes.

Prometaphase I

The key event in prometaphase I is the attachment of the spindle fiber microtubules to the kinetochore proteins at the centromeres. Kinetochore proteins are multiprotein complexes that bind the centromeres of a chromosome to the microtubules of the mitotic spindle. Microtubules grow from centrosomes placed at opposite poles of the cell. The microtubules move toward the middle of the cell and attach to one of the two fused homologous chromosomes. The microtubules attach at each chromosomes' kinetochores. With each member of the homologous pair attached to opposite poles of the cell, in the next phase, the microtubules can pull the homologous pair apart. A spindle fiber that has attached to a kinetochore is called a kinetochore microtubule. At the end of prometaphase I, each tetrad is attached to microtubules from both poles, with one homologous chromosome facing each pole. The homologous chromosomes are still held together at chiasmata. In addition, the nuclear membrane has broken down entirely.

Metaphase I

During metaphase I, the homologous chromosomes are arranged in the center of the cell with the kinetochores facing opposite poles. The homologous pairs orient themselves randomly at the equator. For example, if the two homologous members of chromosome 1 are labeled a and b, then the chromosomes could line up a-b, or b-a. This is important in determining the genes carried by a gamete, as each will only receive one of the two homologous chromosomes. Recall that homologous chromosomes are not identical. They contain slight differences in their genetic information, causing each gamete to have a unique genetic makeup.

This randomness is the physical basis for the creation of the second form of genetic variation in offspring. Consider that the homologous chromosomes of a sexually reproducing organism are originally inherited as two separate sets, one from each parent. Using humans as an example, one set of 23 chromosomes is present in the egg donated by the mother. The father provides the other set of 23 chromosomes in the sperm that fertilizes the egg. Every cell of the multicellular offspring has copies of the original two sets of homologous chromosomes. In prophase I of meiosis, the homologous chromosomes form the tetrads. In metaphase I, these pairs line up at the midway point between the two poles of the cell to form the metaphase plate. Because there is an equal chance that a microtubule fiber will encounter a maternally or paternally inherited chromosome, the arrangement of the tetrads at the metaphase plate is random. Any maternally inherited chromosome may face either pole. Any paternally inherited chromosome may also face either pole. The orientation of each tetrad is independent of the orientation of the other 22 tetrads.

This event—the random (or independent) assortment of homologous chromosomes at the metaphase plate—is the second mechanism that introduces variation into the gametes or spores. In each cell that undergoes meiosis, the arrangement of the tetrads is different. The number of variations is dependent on the number of chromosomes making up a set. There are two possibilities for orientation at the metaphase plate the possible number of alignments therefore equals 2n, where n is the number of chromosomes per set. Humans have 23 chromosome pairs, which results in over eight million (2 23 ) possible genetically-distinct gametes. This number does not include the variability that was previously created in the sister chromatids by crossover. Given these two mechanisms, it is highly unlikely that any two haploid cells resulting from meiosis will have the same genetic composition (Figure 11.4).

To summarize the genetic consequences of meiosis I, the maternal and paternal genes are recombined by crossover events that occur between each homologous pair during prophase I. In addition, the random assortment of tetrads on the metaphase plate produces a unique combination of maternal and paternal chromosomes that will make their way into the gametes.

Anaphase I

In anaphase I, the microtubules pull the linked chromosomes apart. The sister chromatids remain tightly bound together at the centromere. The chiasmata are broken in anaphase I as the microtubules attached to the fused kinetochores pull the homologous chromosomes apart (Figure 11.5).

Telophase I and Cytokinesis

In telophase, the separated chromosomes arrive at opposite poles. The remainder of the typical telophase events may or may not occur, depending on the species. In some organisms, the chromosomes decondense and nuclear envelopes form around the chromatids in telophase I. In other organisms, cytokinesis—the physical separation of the cytoplasmic components into two daughter cells—occurs without reformation of the nuclei. In nearly all species of animals and some fungi, cytokinesis separates the cell contents via a cleavage furrow (constriction of the actin ring that leads to cytoplasmic division). In plants, a cell plate is formed during cell cytokinesis by Golgi vesicles fusing at the metaphase plate. This cell plate will ultimately lead to the formation of cell walls that separate the two daughter cells.

Two haploid cells are the end result of the first meiotic division. The cells are haploid because at each pole, there is just one of each pair of the homologous chromosomes. Therefore, only one full set of the chromosomes is present. This is why the cells are considered haploid—there is only one chromosome set, even though each homolog still consists of two sister chromatids. Recall that sister chromatids are merely duplicates of one of the two homologous chromosomes (except for changes that occurred during crossing over). In meiosis II, these two sister chromatids will separate, creating four haploid daughter cells.

Link to Learning

Review the process of meiosis, observing how chromosomes align and migrate, at Meiosis: An Interactive Animation.

  1. Errors can arise only during the recombination process, which may result in deletions, duplications or translocations causing such abnormalities.
  2. Errors occur when a pair of homologous chromosomes fails to separate during anaphase I or when sister chromatids fail to separate during anaphase II, producing daughter cells with unequal numbers of chromosomes.
  3. Errors occur only during anaphase I of meiosis as chromosomes separate prematurely, triggering aberrations that result in unequal numbers of chromosomes in daughter cells.
  4. Errors during meiosis introduce variations in the DNA sequence that cause changes throughout the phases of meiosis, the intensity of which depend specifically on the size of the variant.

Meiosis II

In some species, cells enter a brief interphase, or interkinesis , before entering meiosis II. Interkinesis lacks an S phase, so chromosomes are not duplicated. The two cells produced in meiosis I go through the events of meiosis II in synchrony. During meiosis II, the sister chromatids within the two daughter cells separate, forming four new haploid gametes. The mechanics of meiosis II is similar to mitosis, except that each dividing cell has only one set of homologous chromosomes. Therefore, each cell has half the number of sister chromatids to separate out as a diploid cell undergoing mitosis.

Prophase II

If the chromosomes decondensed in telophase I, they condense again. If nuclear envelopes were formed, they fragment into vesicles. The centrosomes that were duplicated during interkinesis move away from each other toward opposite poles, and new spindles are formed.

Prometaphase II

The nuclear envelopes are completely broken down, and the spindle is fully formed. Each sister chromatid forms an individual kinetochore that attaches to microtubules from opposite poles.

Metaphase II

The sister chromatids are maximally condensed and aligned at the equator of the cell.

Anaphase II

The sister chromatids are pulled apart by the kinetochore microtubules and move toward opposite poles. Non-kinetochore microtubules elongate the cell.

Telophase II and Cytokinesis

The chromosomes arrive at opposite poles and begin to decondense. Nuclear envelopes form around the chromosomes. Cytokinesis separates the two cells into four unique haploid cells. At this point, the newly formed nuclei are both haploid. The cells produced are genetically unique because of the random assortment of paternal and maternal homologs and because of the recombining of maternal and paternal segments of chromosomes (with their sets of genes) that occurs during crossover. The entire process of meiosis is outlined in Figure 11.6.

Comparing Meiosis and Mitosis

Mitosis and meiosis are both forms of division of the nucleus in eukaryotic cells. They share some similarities, but also exhibit distinct differences that lead to very different outcomes (Figure 11.7). Mitosis is a single nuclear division that results in two nuclei that are usually partitioned into two new cells. The nuclei resulting from a mitotic division are genetically identical to the original nucleus. They have the same number of sets of chromosomes, one set in the case of haploid cells and two sets in the case of diploid cells. In most plants and all animal species, it is typically diploid cells that undergo mitosis to form new diploid cells. In contrast, meiosis consists of two nuclear divisions resulting in four nuclei that are usually partitioned into four new cells. The nuclei resulting from meiosis are not genetically identical and they contain one chromosome set only. This is half the number of chromosome sets in the original cell, which is diploid.

The main differences between mitosis and meiosis occur in meiosis I, which is a very different nuclear division than mitosis. In meiosis I, the homologous chromosome pairs become associated with each other, are bound together with the synaptonemal complex, develop chiasmata and undergo crossover between sister chromatids, and line up along the metaphase plate in tetrads with kinetochore fibers from opposite spindle poles attached to each kinetochore of a homolog in a tetrad. All of these events occur only in meiosis I.

When the chiasmata resolve and the tetrad is broken up with the homologs moving to one pole or another, the ploidy level—the number of sets of chromosomes in each future nucleus—has been reduced from two to one. For this reason, meiosis I is referred to as a reduction division . There is no such reduction in ploidy level during mitosis.

Meiosis II is much more analogous to a mitotic division. In this case, the duplicated chromosomes (only one set of them) line up on the metaphase plate with divided kinetochores attached to kinetochore fibers from opposite poles. During anaphase II, as in mitotic anaphase, the kinetochores divide and one sister chromatid—now referred to as a chromosome—is pulled to one pole while the other sister chromatid is pulled to the other pole. If it were not for the fact that there had been crossover, the two products of each individual meiosis II division would be identical (like in mitosis). Instead, they are different because there has always been at least one crossover per chromosome. Meiosis II is not a reduction division because although there are fewer copies of the genome in the resulting cells, there is still one set of chromosomes, as there was at the end of meiosis I.

Evolution Connection

The Mystery of the Evolution of Meiosis

Some characteristics of organisms are so widespread and fundamental that it is sometimes difficult to remember that they evolved like other simpler traits. Meiosis is such an extraordinarily complex series of cellular events that biologists have had trouble hypothesizing and testing how it may have evolved. Although meiosis is inextricably entwined with sexual reproduction and its advantages and disadvantages, it is important to separate the questions of the evolution of meiosis and the evolution of sex, because early meiosis may have been advantageous for different reasons than it is now. Thinking outside the box and imagining what the early benefits from meiosis might have been is one approach to uncovering how it may have evolved.

Meiosis and mitosis share obvious cellular processes and it makes sense that meiosis evolved from mitosis. The difficulty lies in the clear differences between meiosis I and mitosis. Adam Wilkins and Robin Holliday 1 summarized the unique events that needed to occur for the evolution of meiosis from mitosis. These steps are homologous chromosome pairing, crossover exchanges, sister chromatids remaining attached during anaphase, and suppression of DNA replication in interphase. They argue that the first step is the hardest and most important, and that understanding how it evolved would make the evolutionary process clearer. They suggest genetic experiments that might shed light on the evolution of synapsis.

There are other approaches to understanding the evolution of meiosis in progress. Different forms of meiosis exist in single-celled protists. Some appear to be simpler or more “primitive” forms of meiosis. Comparing the meiotic divisions of different protists may shed light on the evolution of meiosis. Marilee Ramesh and colleagues 2 compared the genes involved in meiosis in protists to understand when and where meiosis might have evolved. Although research is still ongoing, recent scholarship into meiosis in protists suggests that some aspects of meiosis may have evolved later than others. This kind of genetic comparison can tell us what aspects of meiosis are the oldest and what cellular processes they may have borrowed from in earlier cells.


Differences in Purpose

Though both types of cell division are found in many animals, plants, and fungi, mitosis is more common than meiosis and has a wider variety of functions. Not only is mitosis responsible for asexual reproduction in single-celled organisms, but it is also what enables cellular growth and repair in multicellular organisms, such as humans. In mitosis, a cell makes an exact clone of itself. This process is what is behind the growth of children into adults, the healing of cuts and bruises, and even the regrowth of skin, limbs, and appendages in animals like geckos and lizards.

Meiosis is a more specific type of cell division (of germ cells, in particular) that results in gametes, either eggs or sperm, that contain half of the chromosomes found in a parent cell. Unlike mitosis with its many functions, meiosis has a narrow but significant purpose: assisting sexual reproduction. It is the process that enables children to be related but still different from their two parents.

Meiosis and Genetic Diversity

Sexual reproduction uses the process of meiosis to increase genetic diversity. Offspring created through asexual reproduction (mitosis) are genetically identical to their parent, but the germ cells created during meiosis are different from their parent cells. Some mutations frequently occur during meiosis. Further, germ cells have only one set of chromosomes, so two germ cells are required to make a complete set of genetic material for the offspring. The offspring is therefore able to inherit genes from both parents and both sets of grandparents.

Genetic diversity makes a population more resilient and adaptable to the environment, which increases chances of survival and evolution for the long term.

Mitosis as a form of reproduction for single-cell organisms originated with life itself, around 3.8 billion years ago. Meiosis is thought to have appeared around 1.4 billion years ago.


Electronic supplementary material is available online at http://dx.doi.org/10.6084/m9.figshare.c.3825526.v3.

Published by the Royal Society. All rights reserved.

References

. 2011 Using parthenogenetic lineages to identify advantages of sex . Evol. Biol. 38, 115–123. (doi:10.1007/s11692-011-9113-z) Crossref, Google Scholar

. 1978 The evolution of sex . Cambridge, UK : Cambridge University Press . Google Scholar

. 1982 The masterpiece of nature: the evolution and genetics of sexuality . Berkeley, CA : University of California Press . Google Scholar

Neiman M, Sharbel T, Schwander T

. 2014 Genetic causes of transitions from sexual reproduction to asexuality in plants and animals . J. Evol. Biol. 27, 1346–1359. (doi:10.1111/jeb.12357) Crossref, PubMed, Google Scholar

Meirmans S, Meirmans PG, Kirkendall LR

. 2012 The costs of sex: facing real-world complexities . Q. Rev. Biol. 87, 19–40. (doi:10.1086/663945) Crossref, PubMed, ISI, Google Scholar

. 1998 Why sex and recombination? Science 281, 1986–1990. (doi:10.1126/science.281.5385.1986) Crossref, PubMed, ISI, Google Scholar

Lorenzo-Carballa O, Cordero-Rivera A

. 2007 Are parthenogenetic and sexual Ischnura hastata damselflies equally fertile? Testing sexual conflict theories . Ethol. Ecol. Evol. 19, 291–298. (doi:10.1080/08927014.2007.9522552) Crossref, Google Scholar

Schön I, Martens K, van Dijk P

. 2009 Lost sex . Heidelberg, Germany : Springer . Crossref, Google Scholar

. 2004 Developmental success, stability, and plasticity in closely related parthenogenetic and sexual lizards (Heteronotia, Gekkonidae) . Evolution 58, 1560–1572. (doi:10.1111/j.0014-3820.2004.tb01736.x) Crossref, PubMed, Google Scholar

Nagaoka SI, Hassold TJ, Hunt PA

. 2012 Human aneuploidy: mechanisms and new insights into an age-old problem . Nat. Rev. Genet. 13, 493–504. (doi:10.1038/nrg3245) Crossref, PubMed, Google Scholar

. 2009 The evolution of meiosis from mitosis . Genetics 181, 3–12. (doi:10.1534/genetics.108.099762) Crossref, PubMed, Google Scholar

. 2007 On the origin of meiosis in eukaryotic evolution: coevolution of meiosis and mitosis from feeble beginnings . In Recombination and meiosis (eds

), pp. 249–288. Berlin, Germany : Springer . Google Scholar

. 2011 Checkpoint mechanisms: the puppet masters of meiotic prophase . Trends Cell Biol. 21, 393–400. (doi:10.1016/j.tcb.2011.03.004) Crossref, PubMed, Google Scholar

. 2001 To err (meiotically) is human: the genesis of human aneuploidy . Nat. Rev. Genet. 2, 280–291. (doi:10.1038/35066065) Crossref, PubMed, ISI, Google Scholar

. 2010 Genetics of mammalian meiosis: regulation, dynamics and impact on fertility . Nat. Rev. Genet. 11, 124–136. (doi:10.1038/nrg2723) Crossref, PubMed, ISI, Google Scholar

Caryl AP, Jones GH, Franklin FCH

. 2003 Dissecting plant meiosis using Arabidopsis thaliana mutants . J. Exp. Bot. 54, 25–38. (doi:10.1093/jxb/erg041) Crossref, PubMed, Google Scholar

. 1980 Meiosis in Neurospora crassa. I. The isolation of recessive mutants defective in the production of viable ascospores . Genetics 96, 367–378. PubMed, Google Scholar

. 1971 Reproduction and the mechanism of meiotic restitution in the parthenogenetic lizard Cnemidophorus uniparens . J. Morphol. 133, 139–165. (doi:10.1002/jmor.1051330203) Crossref, PubMed, Google Scholar

. 2004 Physiological dependence on copulation in parthenogenetic females can reduce the cost of sex . Anim. Behav. 67, 811–822. (doi:10.1016/j.anbehav.2003.05.014) Crossref, Google Scholar

. 1970 Heterozygosity and genetic polymorphism in parthenogenetic animals . In Essays in evolution and genetics in honor of Theodosius Dobzhansky (eds

), pp. 237–262. Berlin, Germany : Springer . Crossref, Google Scholar

. 2009 A graphical approach to lineage selection between clonals and sexuals . In Lost sex (eds

Schön I, Martens K, van Dijk P

), pp. 75–97. Heidelberg, Germany : Springer . Crossref, Google Scholar

. 1982 The prophecies of parthenogenesis . In Evolution and genetics of life histories (eds

), pp. 75–101. New York , NY : Springer . Google Scholar

. 1954 Parthenogenesis in Drosophila . Genetics 39, 4. PubMed, ISI, Google Scholar

. 1965 Twelve year summary of selection for parthenogenesis in Beltsville small white turkeys . Br. Poult. Sci. 6, 1–6. (doi:10.1080/00071666508415546) Crossref, PubMed, Google Scholar

Cocco J, Butnariu A, Bessa E, Pasini A

. 2013 Sex produces as numerous and long-lived offspring as parthenogenesis in a new parthenogenetic insect . Can. J. Zool. 91, 187–190. (doi:10.1139/cjz-2012-0289) Crossref, Google Scholar

Feldheim KA, Chapman DD, Sweet D, Fitzpatrick S, Prodöhl PA, Shivji MS, Snowden B

. 2010 Shark virgin birth produces multiple, viable offspring . J. Hered. 101, 374–377. (doi:10.1093/jhered/esp129) Crossref, PubMed, Google Scholar

Jokela J, Lively CM, Dybdahl MF, Fox JA

. 1997 Evidence for a cost of sex in the freshwater snail Potamopyrgus antipodarum . Ecology 78, 452–460. (doi:10.1890/0012-9658(1997)078[0452:EFACOS]2.0.CO2) Crossref, Google Scholar

. 1966 The sex of hatchlings of five apparently unisexual species of whiptail lizards (Cnemidophorus, Teiidae) . Am. Midl. Nat. 76, 369–378. (doi:10.2307/2423092) Crossref, Google Scholar

. 1981 The adaptive and evolutionary significance of wing polymorphism and parthenogenesis in Ptinella Motschulsky (Coleoptera: Ptiliidae) . Ecol. Entomol. 6, 89–98. (doi:10.1111/j.1365-2311.1981.tb00975.x) Crossref, Google Scholar

. 2012 Observations on parthenogenesis in monitor lizards . Biawak 6, 11–21. Google Scholar

. 1975 Sex and evolution . Princeton, NJ : Princeton University Press . Google Scholar

2014 The COMPADRE Plant Matrix Database: an open online repository for plant demography . J. Ecol. 103, 202–218. (doi:10.1111/1365-2745.12334) Crossref, Google Scholar

. 1990 Effect of incubation and preservation on resting egg hatching and mixis in the derived clones of the rotifer Brachionus plicatilis . In Rotifer symposium V (eds

), pp. 415–421. Berlin, Germany : Springer . Google Scholar

. 1987 Methods for the activation of the resting eggs of Daphnia . Freshw. Biol. 17, 373–379. (doi:10.1111/j.1365-2427.1987.tb01057.x) Crossref, Google Scholar

. 2013 The two halves of U-shaped mortality . Front. Genet. Aging 4, 31. PubMed, Google Scholar

. 2014 A linear-time algorithm for Gaussian and non-Gaussian trait evolution models . Syst. Biol. 63, 397–408. (doi:10.1093/sysbio/syu005) Crossref, PubMed, ISI, Google Scholar

Freckleton RP, Harvey PH, Pagel M

. 2002 Phylogenetic analysis and comparative data: a test and review of evidence . Am. Nat. 160, 712–726. (doi:10.1086/343873) Crossref, PubMed, ISI, Google Scholar

. 2015 The unsolved challenge to phylogenetic correlation tests for categorical characters . Syst. Biol. 64, 127–136. (doi:10.1093/sysbio/syu070) Crossref, PubMed, ISI, Google Scholar

. 2004 Model selection in ecology and evolution . Trends Ecol. Evol. 19, 101–108. (doi:10.1016/j.tree.2003.10.013) Crossref, PubMed, ISI, Google Scholar

. 1996 Evidence against a frequency-dependent advantage for sexual reproduction in Allium vineale . Am. Nat. 147, 718–734. (doi:10.1086/285876) Crossref, Google Scholar

Kollmann J, Steinger T, Roy BA

. 2000 Evidence of sexuality in European Rubus (Rosaceae) species based on AFLP and allozyme analysis . Am. J. Bot. 87, 1592–1598. (doi:10.2307/2656735) Crossref, PubMed, Google Scholar

2016 An update of the Angiosperm Phylogeny Group classification for the orders and families of flowering plants: APG IV . Bot. J. Linn. Soc. 181, 1–20. (doi:10.1111/boj.12385) Crossref, ISI, Google Scholar

Bradler S, Robertson JA, Whiting MF

. 2014 A molecular phylogeny of Phasmatodea with emphasis on Necrosciinae, the most species-rich subfamily of stick insects . Syst. Entomol. 39, 205–222. (doi:10.1111/syen.12055) Crossref, ISI, Google Scholar

Regier JC, Shultz JW, Zwick A, Hussey A, Ball B, Wetzer R, Martin JW, Cunningham CW

. 2010 Arthropod relationships revealed by phylogenomic analysis of nuclear protein-coding sequences . Nature 463, 1079–1083. (doi:10.1038/nature08742) Crossref, PubMed, ISI, Google Scholar

Pyron RA, Burbrink FT, Wiens JJ

. 2013 A phylogeny and revised classification of Squamata, including 4161 species of lizards and snakes . BMC Evol. Biol. 13, 1. (doi:10.1186/1471-2148-13-93) Crossref, PubMed, ISI, Google Scholar

. 2012 Animal evolution: interrelationships of the living phyla , 3rd edn. Oxford, UK : Oxford University Press . Google Scholar

Paradis E, Claude J, Strimmer K

. 2004 APE: analyses of phylogenetics and evolution in R language . Bioinformatics 20, 289–290. (doi:10.1093/bioinformatics/btg412) Crossref, PubMed, ISI, Google Scholar

Alsos IG, Müller E, Eidesen PB

. 2013 Germinating seeds or bulbils in 87 of 113 tested Arctic species indicate potential for ex situ seed bank storage . Polar Biol. 36, 819–830. (doi:10.1007/s00300-013-1307-7) Crossref, Google Scholar

Barata C, Hontoria F, Amat F

. 1995 Life history, resting egg formation, and hatching may explain the temporal-geographical distribution of Artemia strains in the Mediterranean basin . Hydrobiologia 298, 295–305. (doi:10.1007/BF00033824) Crossref, Google Scholar

. 1958 Etude de la parthénogenèse facultative de Clitumnus extradentatus Br.(Phasmidae) . Ed. Bull. Biol. Fr. Belg. 92, 87–182. Google Scholar

. 1980 Demography of an undergrowth palm in littoral Cameroon . Biotropica 12, 247–255. (doi:10.2307/2387694) Crossref, ISI, Google Scholar

Burke NW, Crean AJ, Bonduriansky R

. 2015 The role of sexual conflict in the evolution of facultative parthenogenesis: a study on the spiny leaf stick insect . Anim. Behav. 101, 117–127. (doi:10.1016/j.anbehav.2014.12.017) Crossref, ISI, Google Scholar

. 1999 Fitness of alternative modes of reproduction: developmental constraints and the evolutionary maintenance of sex . Proc. R. Soc. Lond. B 266, 471–476. (doi:10.1098/rspb.1999.0661) Link, Google Scholar

Darevsky I, Kupriyanova L, Uzzell T

. 1985 Parthenogenesis in reptiles . In Development B (ed.

), pp. 411–526. New York, NY : Wiley-Interscience . Google Scholar

Fernandes MMJ, Vandekerkhove J, Namiotko T

. 2008 Environmental stability and the distribution of the sexes: insights from life history experiments with the geographic parthenogen Eucypris virens (Crustacea: Ostracoda) . Oikos 117, 829–836. (doi:10.1111/j.0030-1299.2008.16557.x) Crossref, Google Scholar

Ferrer M, Durán R, Méndez M, Dorantes A, Dzib G

. 2011 Dinámica poblacional de genets y ramets de Mammillaria gaumeri cactácea endémica de Yucatán . Bol. Soc. Bot. México 89, 83–105. Google Scholar

. 1961 Diapause and parthenogenesis in the eggs of three species of Phasmatodea . Proc. Linn. Soc. New South Wales 86, 268–279. Google Scholar

Hara M, Kanno H, Hirabuki Y, Takehara A

. 2004 Population dynamics of four understorey shrub species in beech forest . J. Veg. Sci. 15, 475–484. (doi:10.1111/j.1654-1103.2004.tb02286.x) Crossref, Google Scholar

. 1999 Fire and population dynamics of woody plants in a neotropical savanna: matrix model projections . Ecology 80, 1354–1369. (doi:10.1890/0012-9658(1999)080[1354:FAPDOW]2.0.CO2) Crossref, Google Scholar

Koltunow A, Johnson SD, Bicknell RA

. 1998 Sexual and apomictic development in Hieracium . Sex Plant Reprod. 11, 213–230. (doi:10.1007/s004970050144) Crossref, Google Scholar

Kramer MG, Templeton AR, Miller KG

. 2002 Evolutionary implications of developmental instability in parthenogenetic Drosophila mercatorum. II. Comparison of two strains with identical genotypes, but different modes of reproduction . Evol. Dev. 4, 234–241. (doi:10.1046/j.1525-142X.2002.02009.x) Crossref, PubMed, Google Scholar

. 1979 Are parthenogenetic and related bisexual insects equal in fertility? Evolution 33, 774–775. (doi:10.1111/j.1558-5646.1979.tb04731.x) Crossref, PubMed, ISI, Google Scholar

Lin CH, Miriti MN, Goodell K

. 2016 Demographic consequences of greater clonal than sexual reproduction in Dicentra canadensis . Ecol. Evol. 6, 3871–3883. (doi:10.1002/ece3.2163) Crossref, PubMed, Google Scholar

Matsuura K, Fujimoto M, Goka K

. 2004 Sexual and asexual colony foundation and the mechanism of facultative parthenogenesis in the termite Reticulitermes speratus (Isoptera, Rhinotermitidae) . Insectes Soc. 51, 325–332. (doi:10.1007/s00040-004-0746-0) Crossref, Google Scholar

. 2013 Coexistence of sperm-dependent asexuals and their sexual hosts: the role of differences in fitness-related traits . Environ. Biol. Fishes 96, 1111–1121. (doi:10.1007/s10641-012-0107-1) Crossref, Google Scholar

. 1971 Parthenogenesis in psocids (Insecta: Psocoptera) . Am. Zool. 11, 327–339. (doi:10.1093/icb/11.2.327) Crossref, Google Scholar

Mondragón D, Durán R, Ramírez I, Valverde T

. 2004 Temporal variation in the demography of the clonal epiphyte Tillandsia brachycaulos (Bromeliaceae) in the Yucatán Peninsula, Mexico . J. Trop. Ecol. 20, 189–200. (doi:10.1017/S0266467403001287) Crossref, Google Scholar

Newton AA, Schnittker RR, Yu Z, Munday SS, Baumann DP, Neaves WB, Baumann P

. 2016 Widespread failure to complete meiosis does not impair fecundity in parthenogenetic whiptail lizards . Development 143, 4486–4494. (doi:10.1242/dev.141283) Crossref, PubMed, Google Scholar

. 2001 Differentiation in reproductive strategy between sexual and asexual populations of Antennaria parlinii (Asteraceae) . Evol. Ecol. Res. 3, 311–330. Google Scholar

. 1974 Reproductive potential of bisexual Pycnoscelus indicus and clones of its parthenogenetic relative, Pycnoscelus surinamensis . Ann. Entomol. Soc. Am. 67, 215–223. (doi:10.1093/aesa/67.2.215) Crossref, Google Scholar

. 2013 Fecundity of parthenogenetic and sexual forms of the flightless Weevil Scepticus insularis (Coleoptera: Curculionidae) with and without effects from mating . Zool. Sci. 30, 906–912. (doi:10.2108/zsj.30.906) Crossref, PubMed, Google Scholar

Verduijn M, Van Dijk PJ, Van Damme J

. 2004 The role of tetraploids in the sexual–asexual cycle in dandelions (Taraxacum) . Heredity 93, 390–398. (doi:10.1038/sj.hdy.6800515) Crossref, PubMed, Google Scholar

Walker MP, Lewis CJ, Whitman DW

. 1999 Effects of males on the fecundity and fertility of female Romalea microptera grasshoppers . J. Orthoptera Res. 8, 277–283. (doi:10.2307/3503444) Crossref, Google Scholar

. 2003 Meristem fate and bulbil formation in Titanotrichum (Gesneriaceae) . Am. J. Bot. 90, 1696–1707. (doi:10.3732/ajb.90.12.1696) Crossref, PubMed, Google Scholar

Weinzierl RP, Schmidt P, Michiels NK

. 1999 High fecundity and low fertility in parthenogenetic planarians . Invertebr. Biol. 118, 87–94. (doi:10.2307/3227051) Crossref, ISI, Google Scholar

Alavi Y, Rooyen A, Elgar MA, Jones TM, Weeks AR

. In press. Novel microsatellite markers suggest the mechanism of parthenogenesis in Extatosoma tiaratum is automixis with terminal fusion . Insect Sci . (doi:10.1111/1744-7917.12373) Google Scholar

. 2007 The origin of Phoxinus eos-neogaeus unisexual hybrids . Mol. Ecol. 16, 4562–4571. (doi:10.1111/j.1365-294X.2007.03511.x) Crossref, PubMed, Google Scholar

. 1986 Allozyme divergence among five diploid species of Antennaria (Asteraceae: Inuleae) and their allopolyploid derivatives . Am. J. Bot . 73, 287–296. (doi:10.2307/2444183) Crossref, Google Scholar

2010 Exceptional cryptic diversity and multiple origins of parthenogenesis in a freshwater ostracod . Mol. Phylogen. Evol. 54, 542–552. (doi:10.1016/j.ympev.2009.08.022) Crossref, PubMed, Google Scholar

Brochmann C, Xiang Q, Brunsfeld S, Soltis D, Soltis P

. 1998 Molecular evidence for polyploid origins in Saxifraga (Saxifragaceae): the narrow arctic endemic S. svalbardensis and its widespread allies . Am. J. Bot. 85, 135. (doi:10.2307/2446562) Crossref, PubMed, Google Scholar

. 1990 Speciation by hybridization in phasmids and other insects . Can. J. Zool. 68, 1747–1760. (doi:10.1139/z90-256) Crossref, Google Scholar

Cole CJ, Dessauer HC, Barrowclough GF

. 1988 Hybrid origin of a unisexual species of whiptail lizard, Cnemidophorus neomexicanus, in western North America: new evidence and a review . Am. Mus. Novit. 2905, 1–38. Google Scholar

Conti E, Soltis DE, Hardig TM, Schneider J

. 1999 Phylogenetic relationships of the silver saxifrages (Saxifraga, sect. Ligulatae Haworth): implications for the evolution of substrate specificity, life histories, and biogeography . Mol. Phylogenet. Evol. 13, 536–555. (doi:10.1006/mpev.1999.0673) Crossref, PubMed, Google Scholar

Corley L, Blankenship J, Moore A

. 2001 Genetic variation and asexual reproduction in the facultatively parthenogenetic cockroach Nauphoeta cinerea: implications for the evolution of sex . J. Evol. Biol. 14, 68–74. (doi:10.1046/j.1420-9101.2001.00254.x) Crossref, PubMed, Google Scholar

Corley LS, Blankenship JR, Moore AJ, Moore PJ

. 1999 Developmental constraints on the mode of reproduction in the facultatively parthenogenetic cockroach Nauphoeta cinerea . Evol. Dev. 1, 90–99. (doi:10.1046/j.1525-142x.1999.99001.x) Crossref, PubMed, Google Scholar

. 1972 Chromosomal diversity in the Australian Phasmatodea . Aust. J. Zool. 20, 445–462. (doi:10.1071/ZO9720445) Crossref, Google Scholar

Gillespie LJ, Archambault A, Soreng RJ

. 2007 Phylogeny of Poa (Poaceae) based on trnT–trnF sequence data: major clades and basal relationships . Aliso: J. Syst. Evol. Bot. 23, 420–434. Google Scholar

Hand M, Vít P, Krahulcová A, Johnson S, Oelkers K, Siddons H, Chrtek J, Fehrer J, Koltunow A

. 2015 Evolution of apomixis loci in Pilosella and Hieracium (Asteraceae) inferred from the conservation of apomixis-linked markers in natural and experimental populations . Heredity 114, 17–26. (doi:10.1038/hdy.2014.61) Crossref, PubMed, Google Scholar

Kearney M, Fujita MK, Ridenour J

. 2009 Lost sex in the reptiles: constraints and correlations . In Lost sex (eds

Schön I, Martens K, van Dijk P

), pp. 447–474. Heidelberg, Germany : Springer . Crossref, Google Scholar

. 2001 Life-history changes that accompany the transition from sexual to parthenogenetic reproduction in Drosophila mercatorum . Evolution 55, 748–761. (doi:10.1554/0014-3820(2001)055[0748:LHCTAT]2.0.CO2) Crossref, PubMed, ISI, Google Scholar

. 1987 Cytological mechanisms of thelytokous parthenogenesis in insects . Genome 29, 367–369. (doi:10.1139/g87-062) Crossref, Google Scholar

Lorenzo-Carballa M, Cordero-Rivera A

. 2009 Thelytokous parthenogenesis in the damselfly Ischnura hastata (Odonata, Coenagrionidae): genetic mechanisms and lack of bacterial infection . Heredity 103, 377–384. (doi:10.1038/hdy.2009.65) Crossref, PubMed, Google Scholar

Lorenzo-Carballa M, Hadrys H, Cordero-Rivera A, Andrés J

. 2012 Population genetic structure of sexual and parthenogenetic damselflies inferred from mitochondrial and nuclear markers . Heredity 108, 386–395. (doi:10.1038/hdy.2011.84) Crossref, PubMed, Google Scholar

Lorenzo-Carballa MO, Hassall C, Encalada AC, Sanmartín-Villar I, Torres-Cambas Y, Cordero-Rivera A

. 2016 Parthenogenesis did not consistently evolve in insular populations of Ischnura hastata (Odonata, Coenagrionidae) . Ecol. Entomol . 42, 67–76. (doi:10.1111/een.12360) Crossref, Google Scholar

. 2013 Origin and genetic diversity of diploid parthenogenetic Artemia in Eurasia . PLoS ONE 8, e83348. (doi:10.1371/journal.pone.0083348) Crossref, PubMed, Google Scholar

. 1993 The origin and evolution of parthenogenesis in the Heteronotia binoei complex: synthesis . Genetica 90, 269–280. (doi:10.1007/BF01435044) Crossref, Google Scholar

Muñoz J, Gómez A, Green AJ, Figuerola J, Amat F, Rico C

. 2010 Evolutionary origin and phylogeography of the diploid obligate parthenogen Artemia parthenogenetica (Branchiopoda: Anostraca) . PLoS ONE 5, e11932. (doi:10.1371/journal.pone.0011932) Crossref, PubMed, Google Scholar

. 1984 Reaction norms of development rate among diploid clones of the parthenogenetic cockroach Pycnoscelus surinamensis . Evolution 38, 1186–1193. (doi:10.1111/j.1558-5646.1984.tb05642.x) Crossref, PubMed, Google Scholar

Pongratz N, Storhas M, Carranza S, Michiels NK

. 2003 Phylogeography of competing sexual and parthenogenetic forms of a freshwater flatworm: patterns and explanations . BMC Evol. Biol. 3, 23. (doi:10.1186/1471-2148-3-23) Crossref, PubMed, ISI, Google Scholar

. 1973 The origin of Taraxacum agamospecies . Bot. J. Linn. Soc. 66, 189–211. (doi:10.1111/j.1095-8339.1973.tb02169.x) Crossref, ISI, Google Scholar

Rossi V, Piotti A, Baltanás A, Benassi G, Menozzi P

. 2008 Genetic diversity and mixed reproduction in Eucypris virens (Crustacea: Ostracoda) . Fund. Appl. Limnol./Arch. Hydrobiol. 172, 147–159. (doi:10.1127/1863-9135/2008/0172-0147) Crossref, Google Scholar

. 1968 Chromosomes of the Pycnoscelus indicus and P. surinamensis complex (Blattaria: Blaberidae: Pycnoscelinae) . Psyche 75, 53–76. (doi:10.1155/1968/38048) Crossref, Google Scholar

Ryabinina N, Grechko V, Semenova S, Darevsky IS

. 2011 On the hybridogenous origin of the parthenogenetic species Lacerta dahli and Lacerta rostombekovi revealed by RAPD technique . Russ. J. Herpetol. 6, 55–60. Google Scholar

Xu S, Innes DJ, Lynch M, Cristescu ME

. 2013 The role of hybridization in the origin and spread of asexuality in Daphnia . Mol. Ecol. 22, 4549–4561. (doi:10.1111/mec.12407) Crossref, PubMed, Google Scholar

van Baarlen P, van Dijk PJ, Hoekstra RF, de Jong JH

. 2000 Meiotic recombination in sexual diploid and apomictic triploid dandelions (Taraxacum officinale L.) . Genome 43, 827–835. (doi:10.1139/g00-047) Crossref, PubMed, ISI, Google Scholar

Baxevanis AD, Kappas I, Abatzopoulos TJ

. 2006 Molecular phylogenetics and asexuality in the brine shrimp Artemia . Mol. Phylogenet. Evol. 3, 724–738. (doi:10.1016/j.ympev.2006.04.010) Crossref, Google Scholar

. 1973 Fetal mortality due to aneuploidy and irregular meiotic segregation in the mouse . In Proc. symp. Institut National de la Santé et de la Recherche Medicale (eds

), pp. 255–268. Paris, France : Institut National de la Santé et de la Recherche Medicale . Google Scholar

Kuliev A, Zlatopolsky Z, Kirillova I, Spivakova J, Janzen JC

. 2011 Meiosis errors in over 20,000 oocytes studied in the practice of preimplantation aneuploidy testing . Reprod. Biomed. Online 22, 2–8. (doi:10.1016/j.rbmo.2010.08.014) Crossref, PubMed, Google Scholar

Ni M, Feretzaki M, Li W, Floyd-Averette A, Mieczkowski P, Dietrich FS, Heitman J

. 2013 Unisexual and heterosexual meiotic reproduction generate aneuploidy and phenotypic diversity de novo in the yeast Cryptococcus neoformans . PLoS Biol. 11, e1001653. (doi:10.1371/journal.pbio.1001653) Crossref, PubMed, Google Scholar

Suomalainen E, Saura A, Lokki J

. 2012 Parthenogenetic insects . In Evolutionary biology (ed.

), pp. 209–250. Berlin, Germany : Springer . Google Scholar

Parker ED, Selander RK, Hudson RO, Lester L

. 1977 Genetic diversity in colonizing parthenogenetic cockroaches . Evolution 31, 836–842. (doi:10.1111/j.1558-5646.1977.tb01076.x) Crossref, PubMed, ISI, Google Scholar

. 1996 Parasexual recombination in fungi . J. Genet. 75, 281–286. (doi:10.1007/BF02966308) Crossref, Google Scholar

. 1983 Interruption of synthesis as a cost of sex in small organisms . Am. Nat. 121, 825–833. (doi:10.1086/284106) Crossref, Google Scholar

. 2004 Morphological and physiological correlates of hybrid parthenogenesis . Am. Nat. 164, 803–813. (doi:10.1086/425986) Crossref, PubMed, Google Scholar

Wilmut I, Sales D, Ashworth C

. 1986 Maternal and embryonic factors associated with prenatal loss in mammals . J. Reprod. Fertil. 76, 851–864. (doi:10.1530/jrf.0.0760851) Crossref, PubMed, Google Scholar

. 2011 Before senescence: the evolutionary demography of ontogenesis . Proc. R. Soc. B 278, 801–809. (doi:10.1098/rspb.2010.2190) Link, Google Scholar


Examples of Haploid Cells

Almost all of the cells in sexually reproducing organisms are diploid, with the only exception being the gametes. Sperm and egg cells each contain a single set of chromosomes which, during sexual reproduction, combine to form a single diploid cell (or zygote). Examples of haploid gametes include:

  • Sperm and egg cells (the reproductive cells of humans)
  • Spores (the reproductive cells of fungi, algae, and plants) (the reproductive cells of male plants)


Meiosis and genetic variation - PowerPoint PPT Presentation

PowerShow.com is a leading presentation/slideshow sharing website. Whether your application is business, how-to, education, medicine, school, church, sales, marketing, online training or just for fun, PowerShow.com is a great resource. And, best of all, most of its cool features are free and easy to use.

You can use PowerShow.com to find and download example online PowerPoint ppt presentations on just about any topic you can imagine so you can learn how to improve your own slides and presentations for free. Or use it to find and download high-quality how-to PowerPoint ppt presentations with illustrated or animated slides that will teach you how to do something new, also for free. Or use it to upload your own PowerPoint slides so you can share them with your teachers, class, students, bosses, employees, customers, potential investors or the world. Or use it to create really cool photo slideshows - with 2D and 3D transitions, animation, and your choice of music - that you can share with your Facebook friends or Google+ circles. That's all free as well!

For a small fee you can get the industry's best online privacy or publicly promote your presentations and slide shows with top rankings. But aside from that it's free. We'll even convert your presentations and slide shows into the universal Flash format with all their original multimedia glory, including animation, 2D and 3D transition effects, embedded music or other audio, or even video embedded in slides. All for free. Most of the presentations and slideshows on PowerShow.com are free to view, many are even free to download. (You can choose whether to allow people to download your original PowerPoint presentations and photo slideshows for a fee or free or not at all.) Check out PowerShow.com today - for FREE. There is truly something for everyone!

presentations for free. Or use it to find and download high-quality how-to PowerPoint ppt presentations with illustrated or animated slides that will teach you how to do something new, also for free. Or use it to upload your own PowerPoint slides so you can share them with your teachers, class, students, bosses, employees, customers, potential investors or the world. Or use it to create really cool photo slideshows - with 2D and 3D transitions, animation, and your choice of music - that you can share with your Facebook friends or Google+ circles. That's all free as well!


Abstract

Meiosis is a key event of sexual life cycles in eukaryotes. Its mechanistic details have been uncovered in several model organisms, and most of its essential features have received various and often contradictory evolutionary interpretations. In this perspective, we present an overview of these often ‘weird’ features. We discuss the origin of meiosis (origin of ploidy reduction and recombination, two-step meiosis), its secondary modifications (in polyploids or asexuals, inverted meiosis), its importance in punctuating life cycles (meiotic arrests, epigenetic resetting, meiotic asymmetry, meiotic fairness) and features associated with recombination (disjunction constraints, heterochiasmy, crossover interference and hotspots). We present the various evolutionary scenarios and selective pressures that have been proposed to account for these features, and we highlight that their evolutionary significance often remains largely mysterious. Resolving these mysteries will likely provide decisive steps towards understanding why sex and recombination are found in the majority of eukaryotes.

This article is part of the themed issue ‘Weird sex: the underappreciated diversity of sexual reproduction’.

1. Introduction

In eukaryotic sexual life cycles, haploid cells fuse to give rise to diploids, before diploid cells are converted back to haploids in a process known as meiosis. Meiosis reduces a cell's chromosome number by half, while also creating new allele combinations distributed across daughter cells through segregation and recombination. This genetic reshuffling reduces genetic associations within and between loci and is thought to be the basis of the success of sexual reproduction. Mechanistic studies of meiosis have been carried out in different fields, such as cell biology, genetics and epigenetics, encompassing a wide range of eukaryotes. However, these studies rarely focus on the evolutionary significance of meiotic mechanisms, rather mentioning them in passing and often in a simplified manner. In evolutionary biology studies, meiosis is often simplified and represented by random assortment of chromosomes and recombination maps expressing the probability of recombination events between ordered loci, with little attention to the molecular and cellular details. While these simplifications are legitimate and useful in many cases, the wealth of mechanistic findings being uncovered points to a considerable number of evolutionary puzzles surrounding meiosis that have yet to be resolved. Indeed, in the following perspective, we will show that close scrutiny of almost every aspect of meiosis will reveal ‘weird’ features that constitute evolutionary mysteries.

2. The origins of meiosis

The origin of meiosis through gradual steps is among the most intriguing evolutionary enigmas [1,2]. Meiosis is one of the ‘major innovations’ of eukaryotes that evolved before their subsequent radiation over 1 billion years ago [3–5]. Extant eukaryotes share a set of genes specifically associated with meiosis, implying that it evolved only once before their last common ancestor [6,7]. Identifying the selective scenario that led to its early evolution is difficult, but clues can be obtained by determining (i) which mitotic cellular processes were reused in meiosis (e.g. DNA repair through homologous recombination and possibly reduction), (ii) which selective steps were involved in the assembly of the full cellular process, and (iii) why different forms of meiosis were perhaps less successful.

(a) The origin of ploidy reduction

A form of reductional cell division (aka ‘proto-meiosis’) probably evolved in early asexual unicellular eukaryotes. Two scenarios for this have been proposed. The first is that diploidy accidentally occurred by replication of the nuclear genome without subsequent cell division (endoreplication) [8–12], and that returning to haploidy was selected for to correct this. Because either haploidy or higher ploidy levels may be favoured in different ecological situations [13,14], a variant of this scenario is that a proto-meiosis–endoreplication cycle evolved to switch between ploidy levels [5]. The resulting life cycle may have resembled modern ‘parasexual’ fungi in which diploid cells lose chromosomes in subsequent mitotic divisions, leading to haploidy via aneuploid intermediates [15]. Many other modern eukaryotes also increase and decrease their ploidy somatically, depending on growth stage or specific environmental stimuli [16]. The second scenario is that proto-meiosis evolved in response to the fusion of two haploid cells (syngamy), as in standard modern eukaryotic sexual life cycles. Syngamy may have been favoured because it allows recessive deleterious mutations to be masked in diploids [1,12]. A difficulty with this idea is that such masking may not be sufficient to favour diploidy in asexuals [17]. In a variant of this scenario, early syngamy evolved as a result of ‘manipulation’ by selfish elements (plasmids, transposons) to promote their horizontal transmission [18]. In support of this view, mating-type switching (which can allow syngamy in haploid colonies) has evolved multiple times in yeasts and involves domesticated mobile genetic elements [19].

(b) The origin of homologue pairing and meiotic recombination

Meiosis requires the correct segregation of homologues, which is achieved by homologue pairing at the beginning of prophase I (figure 1). This homology search is mediated by the active formation of numerous DNA double-strand breaks (DSBs) followed by chiasmata formation, but less well-known mechanisms of recombination-independent pairing also exist [20]. Non-homologous centromere coupling is also often observed at this stage, but the functional and evolutionary significance of this coupling is elusive [21]. In many species, chromosome pairing is further strengthened by ‘synapsis’, which is the formation of a protein structure known as the synaptonemal complex [22] and the pairing of homologous centromeres [21]. Chiasmata are then resolved as either crossovers (hereafter ‘COs’) resulting in the exchange of large chromatid segments, or non-crossovers (NCOs), where both situations cause gene-conversion events [23]. The synaptonemal complex then disappears, and homologues remain tethered at CO positions and centromeres. The precise function of the synaptonemal complex is not entirely understood [20] one possibility is that it may serve to stabilize homologues during CO maturation. Some pairing mechanism must be advantageous to ensure proper segregation of homologues, but the origins and selective advantage of extensive pairing, synapsis, gene conversion and recombination remain poorly understood [24].

Figure 1. Schematic of the different steps in standard meiosis. The top panel illustrates the different phases of a typical female meiosis for each of the two meiotic divisions: prophase (P, with early and late prophase distinguished), metaphase (M), anaphase (A) and telophase (T). The nuclear membrane is indicated by the green contour (dashed when it starts fragmenting). The small black circles represent microtubule organizing centres and the black lines represent microtubules of the meiotic spindle. First and second polar bodies are shown as grey circles next to the oocyte (chromosomes inside the polar bodies are not shown). Homologous chromosomes are represented with the same colour with slightly different shades (e.g. orange and light orange). Homologues pair and segregate in meiosis I, then sister chromatids segregate in meiosis II. The middle panel shows the meiotic cell cycle. The timing of the primary meiotic arrest is indicated by a red star, while the timing of the most common secondary arrests in different organisms is indicated by green stars (see §4a). The lower panel indicates the important steps (DSB formation, crossing overs) occurring during prophase I. The synaptonemal complex is shown in yellow. Chromatin condenses in chromosomes throughout prophase I (only one pair of homologues is illustrated). In most species, telomeres attach to the nuclear envelope. The attachment plate is indicated by a grey bar. MSCI, meiotic sex chromosome inactivation (see §4d).

Most evidence suggests that homologous recombination evolved long before meiosis, as it occurs in all domains of life and involves proteins that share strong homology [25,26]. One hypothesis is that meiotic pairing and extensive homologous recombination in meiosis evolved to avoid the burden and consequences of non-allelic ectopic recombination in the large genomes of early eukaryotes, which presumably had many repetitive sequences [9,27,28]. Such sequences may have been related to the spread of retrotransposons in early eukaryotes, of which many types are very ancient in eukaryotes, but absent in bacteria and archaea [29]. A second possibility is that recombination arose by the spread of self-promoting genetic elements exploiting the machinery of DNA repair and associated gene conversion [30]. Another hypothesis is that pairing and recombination initially arose as a way to repair mutational damage caused by increased oxidative stress due to rising atmospheric oxygen or endosymbiosis [7,31–33]. This scenario presupposes that DNA maintenance is inefficient in the absence of meiosis however, prokaryotes (including archaea) have efficient repair mechanisms that involve recombination, but not meiosis [9]. In addition, this scenario does not fit well with the observation that a large number of DSBs are actively generated at the onset of meiosis [1,34].

(c) The origin of two-step meiosis

A particular feature of meiosis is that it starts with chromosome doubling (S phase figure 1) before meiosis occurs (figure 2a). For ploidy reduction, the initial steps appear superfluous [35]. A simpler single-step cell division, without the initial DNA replication phase, could in principle achieve ploidy reduction (figure 2b). Recombination may not be a crucial difference between one- and two-step meiosis, as both can involve COs, even if with one CO, the two meiotic products carry recombinant chromosomes in one-step meiosis, whereas only two out of four are recombinant in two-step meiosis [36]. Three hypotheses have been proposed to account for two-step meiosis. The first postulates that two-step meiosis better protects against particular selfish genetic elements (SGEs) that increase their transmission frequencies by sabotaging the meiotic products in which they do not end up (known as ‘sister killers’, distinct from the ‘sperm killers’ discussed below) [37]. In a two-step meiosis, there is uncertainty as to whether the reductional division is meiosis I or II, meaning that the sabotage mechanism has a much reduced efficacy. Microsporidia and red algae show specific modifications to meiosis that increase such uncertainty even more [38]. However, such sister killers are hypothetical, and theoretical studies based on assumptions about how different killers might act suggest that this mechanism does not inevitably promote the development of a two-step meiosis [39]. The second hypothesis is that sexual species with one-step meiosis would be vulnerable to invasion by asexual mutants, and have thus gone disproportionally extinct in the past. Contrary to one-step meiosis, most automictic modifications of two-step meiosis involve a loss of heterozygosity with each generation (see §3c), which would cause expression of recessive and partially recessive deleterious mutational effects, and reduce the fitness of newly emerging asexual mutants [36]. Finally, a third hypothesis posits that a one-step meiosis is more complex and thus less likely to evolve than a two-step meiosis [9]. Mitotic and meiotic cell cycles start similarly with DNA replication in response to increasing cyclin-dependent kinase (CDK) activity. Two-step meiosis can be achieved simply by modulating CDK activity at the end of a cell cycle to add a second division event [40]. By contrast, a one-step meiosis would require extensive modification of the mitotic cycle. Despite earlier suggestions of its presence in some basal eukaryotes (protists) [8,41], there are presently no firm indications that one-step meioses exist in nature [38,42], although inverted meiosis (see below) is genetically similar to mitosis followed by single-step meiosis.

Figure 2. Schematic of meiosis and some of its modifications. (a) Regular meiosis. Following DNA replication, homologous chromosomes are separated in the first meiotic division, whereas sister chromatids are separated in the second division. COs result in chromosomes in the final meiotic products that carry genetic material from both homologous chromosomes. (b) Hypothetical ‘one-step’ meiosis, in which DNA replication before entering meiosis is suppressed and, therefore, only a single meiotic division is required. (c) Multivalent formation in a neo-tetraploid. Blue and orange chromosome pairs are assumed to be identical or very similar so that pairing can occur. Chiasmata of one chromosome with three other chromosomes leads to mis-segregation. (d) Bivalent formation in a tetraploid with exactly one CO per chromosome. Chromosomes may pair randomly (leading to polysomic inheritance), but segregation proceeds normally. (e) Inverted meiosis, in which sister chromatids are separated in the first division and homologous chromosomes in the second division. Note that although centromeres are shown here for clarity, all described species consistently using inverted meiosis are holokinetic (no centromeres). (f) Central fusion automixis, a mechanism of producing diploid eggs that can then develop parthenogenetically without fertilization. As a consequence of COs, heterozygosity may be lost with this mechanism in regions distal to the centromere.

3. Secondary modifications of meiosis

Meiosis is remarkably conserved across eukaryotes. Nevertheless, in many species, variants exist that may offer insights into the evolutionary origins and mechanistic constraints of meiosis. Here, we discuss three of these modifications: meiosis in polyploids, inverted meiosis and meiosis in asexual organisms.

(a) Meiosis and polyploidy

Polyploidy is surprisingly common in eukaryotes given the considerable problems it poses to meiosis [43–45]. In diploids, homologous chromosomes recognize each other and align to form bivalents during Prophase I, but when there are three or more chromosomes with sufficient homology, these chromosomes may all align to varying degrees, forming multivalents. This can occur when all chromosome sets originate from the same species (autopolyploidy), and also when polyploidy is a result of hybridization (allopolyploidy). Multivalent formation is often associated with mis-segregation of chromosomes (figure 2c) as well as chromosomal rearrangements arising from recombination within multivalents, leading to reduced fertility and low-fitness offspring (e.g. [46–48]). These problems may be compounded in allopolyploids because recombination homogenises partially differentiated chromosomes, thereby further increasing the likelihood that they will pair (the ‘polyploid ratchet’ [49]).

Given these detrimental effects, the existence of successful polyploid species and lineages indicates that natural selection can often promote transitions from multi- to bivalents that will then segregate as in diploids (cf. figure 2c,d) (e.g. [50,51]). However, how such transitions are achieved at the molecular level remains a mystery. Part of the answer seems to be a reduction in the number of COs, since multivalents can only form with at least two COs per chromosome [51–53]. This mechanism seems particularly important in autopolyploids and may be achieved through increasing CO interference (see §5b for definition) [54]. Several candidate genes that may effect such modifications have been identified in the autotetraploid Arabidopsis arenosa [51,55]. In allopolyploids, there is evidence for genes that have been selected to strengthen the preferential pairing of homologous (i.e. of the same origin, rather than ‘homeologous’) chromosomes, including ph1 in hexaploid wheat [56]. This preferential pairing can also be achieved through reducing CO numbers, but specifically those between homeologues this could indirectly produce an increase in CO numbers and hence recombination rates between homologues [43]. Intriguingly, because most extant organisms have a history of polyploidy, many features of ‘standard’ meiosis such as CO interference may have been shaped by the problems involved in multivalent segregation.

Polyploidy with odd numbers of chromosome sets poses an even greater problem because aneuploid gametes are generally produced (e.g. [57]). However, there are some plant species where solutions to even this problem have evolved, and where odd-number polyploidy appears to persist in a stable manner. In these species, the problem of unequal segregation during meiosis is solved through exclusion of univalents in one sex but inclusion in the other, leading, for example, to haploid sperm and tetraploid eggs in pentaploid dog roses [58].

(b) Inverted meiosis

In normal meiosis, homologous chromosomes are separated during meiotic division I, whereas sister chromatids are separated during meiosis II. Why meiosis generally follows this order is unknown, but interestingly, in some species meiosis takes place in the reverse order (figure 2e), including some flowering plants [59–61], mites [62], true bugs [63] and mealybugs [64]. All species with this ‘inverted’ meiosis described to date seem to have holocentric chromosomes (i.e. the kinetochores are assembled along the entire chromosome, rather than at localized centromeres). Inverted meiosis is viewed as a possible solution to specific problems of kinetochore geometry in such meiosis [65]. Yet, intriguing as they are, these systems provide little insight into why inverted meiosis is absent or very rare in monocentric species.

It is conceivable that a reverse order of divisions would make meiosis more vulnerable to exploitation by meiotic drive or sister killer SGEs, but to the best of our knowledge, there is currently neither theoretical nor empirical support for this idea. Another possibility is that meiosis I tends to be reductional because it allows for DSB repair by sister chromatid exchange in arrested female meiosis [66]. Alternatively, the order of meiotic divisions could merely be a ‘frozen accident’, i.e. a solution that has been arrived at a long time ago by chance, and that reversal is difficult (at least with monocentric chromosomes). However, a recent paper investigating human female meioses in unprecedented detail casts doubt on this view [67]. The careful genotyping of eggs (or embryos) and polar bodies at many markers indicated that surprisingly often, chromosomes followed an ‘inverted meiosis’ pattern of segregation, even though this led to aneuploidies in approx. 23% of cases. The question of why one order of meiotic divisions is almost universal, therefore, remains unresolved.

(c) Meiosis modifications and loss of sex

Many organisms have abandoned canonical sexual reproduction, reproducing asexually by suppressing or modifying meiosis and producing diploid eggs that can develop without fertilization. This raises two connected mysteries: why are some types of modifications much more frequent than others, and how can mitotic (or mitosis-like) asexual reproduction (‘apomixis’ or ‘clonal parthenogenesis’ in animals, ‘mitotic apomixis’ in plants) evolve from meiosis? Examples of meiosis-derived modes of asexual reproduction include chromosome doubling prior to meiosis (‘endomitosis’ or ‘pre-meiotic doubling’), fusion of two of the four products of a single meiosis (‘automixis’ in animals, ‘within-tetrad mating’ in fungi), and suppression of one of the two meiotic divisions (included under ‘automixis’ or ‘meiotic apomixis’, depending on the author see [68–74] for detailed descriptions of these processes).

Two particularly common modes of asexuality are the suppression of meiosis I, and automixis involving fusing meiotic products that were separated during meiosis I (‘central fusion’, figure 2f). Both are genetically equivalent and lead to reduced heterozygosity when there is recombination between a locus and the centromere of the chromosome on which it is located. Most other forms of meiosis-derived asexual reproduction lead to a much stronger reduction in offspring heterozygosity [75–79], and it has been hypothesized that the reduced fitness of homozygous progeny explains the rarity of these other forms [71,78,80]. Indirect support comes from the observation that species with regular asexual reproduction usually do so by central fusion or suppression of meiosis I, often accompanied by very low levels of recombination, thus maintaining heterozygosity. By contrast, species that only rarely reproduce asexually show a wider variety of asexual modes and higher levels of recombination [1,71,73,81,82]. Nonetheless, this hypothesis cannot explain some observations, for instance, the rarity of pre-meiotic doubling with sister-chromosome pairing, which would also efficiently maintain heterozygosity [71]. Perhaps, evolving a mechanism that ensures exclusive sister pairing (i.e. the complete absence of non-sister pairing) is difficult, though it seems to occur in some lizard species [83]. In addition, such a system would make it difficult to repair DSBs occurring before doubling (as both sister chromatids would have the same DSBs) [71].

The question of how a mitotic asexual mutant can invade a sexual species is at the heart of the debate on the evolutionary maintenance of sex, as this is what is investigated in most theoretical models and is the situation where the cost of sex is most evident [1]. However, unless meiosis can be entirely bypassed (e.g. as with vegetative reproduction), secondary asexuality is likely to evolve via modification of meiosis, keeping much of the cell signalling and machinery intact ([65,76,80,81], see also §4). Indeed, detailed cytological and genetic investigations in several asexual species thought to reproduce clonally by mitotic apomixis have uncovered remnants of meiosis [73,84–86]. In Daphnia, meiosis I is aborted mid-way and a normal meiosis II follows. Hence, clonality in Daphnia is meiotically derived [84]. This should lead to loss of heterozygosity in centromere-distal regions, but if recombination is fully suppressed the genetic outcome resembles mitosis. Importantly, this suggests a possible stepwise route to evolution of mitosis-like asexuality. Rare automixis (spontaneous development of unfertilized eggs) occurs in many species [1,81]. If this becomes more common, forms of automixis maintaining heterozygosity in centromere regions might be selectively favoured and recombination suppressed, eventually leading to meiosis-derived asexuality with the same genetic consequences as mitosis [87–91]. Indeed, in Arabidopsis, meiosis can be transformed to genetically resemble mitosis, but modification of several genes is needed to achieve this [92–94]. In angiosperms, there is also the difficulty to overcome the absence of endosperm fertilization to achieve proper seed development, which further stresses that meiosis-derived asexuality is unlikely to evolve in a single step. To fully understand the evolutionary maintenance of sex, we may therefore need to understand the selection pressures acting in the intermediate stages, which probably involve loss of heterozygosity, and thus inbreeding depression [77,80]. In many cases, the initial evolution of asexuality may thus resemble the evolution of self-fertilization, and several traits may pre-exist (such as low recombination rates) that make the successful transition to asexuality more likely in some taxa.

4. Meiosis punctuates life cycles

Meiosis is a key step in sexual life cycles, as well as some asexual life cycles derived from sexual ancestors. In multicellular eukaryotes, where meiosis is tightly associated with reproduction (unlike in many protists), meiosis is also a cellular and genetic bottleneck at the critical transition between the diploid and the haploid phases.

(a) Meiosis timing and arrest

In early haploid eukaryotes, meiosis probably quickly followed endomitosis or syngamy. Today, multicellular eukaryotes exhibit a variety of life cycles in which the haploid or diploid phase may predominate. The duration of the different phases was perhaps initially controlled, in part, by the timing of meiosis—for instance, a multicellular, extended diploid phase likely evolved by postponing meiosis. However, in metazoans, life cycles are mostly determined by the extent of somatic development within each phase rather than by the timing of meiosis, which can be halted or postponed. In animals, where haploid mitosis is suppressed, syngamy immediately follows meiosis. Furthermore, specific cells are ‘destined’ at an early stage to eventually undergo meiosis (aka germline), whereas this cell fate is determined much later in fungi, plants and some algae.

The timing of meiosis in the germline of animals has been intensively investigated. Whereas male meiosis occurs continuously, female meiosis usually stops twice (figure 1). These ‘meiotic arrests’ are under the control of various factors that are not completely identified across animals [95–97]. Arrest 1 occurs in prophase I during early development and can last years until sexual maturity. The timing of arrest 2 is more variable (ranging from metaphase I in many invertebrates, to metaphase II in vertebrates and G1 phase after meiosis II in some echinoderms), and may have evolved to prevent the risk of premature parthenogenetic cleavage of oocytes or inappropriate DNA replication before fertilization [97,98] this is supported by the fact that this arrest is usually released by fertilization. However, the evolutionary significance of its precise timing in diverse groups is not well understood. Three ideas have been put forward to explain arrest 1 [66]. First, its occurrence at prophase I may allow the repair of accidental DSBs by sister chromatid exchange during long periods between arrests 1 and 2. Second, if arrest 1 was to occur during an earlier mitotic division within the germline, this might decrease the variance in the number of deleterious mutations among gametes within individuals, which may be detrimental if some defective gametes or early embryos can be eliminated and replaced during reproduction. Third, it may be easier to prevent uncontrolled proliferation in a non-dividing meiotic oocyte, as once the cell starts the meiotic cell division, it cannot engage in further mitotic divisions. Arrest 1 may thus have evolved to control (and minimize) the number of possibly wasteful and mutagenic mitotic divisions in the female germline. Similar meiotic arrests in plants are unknown. Plants seem to completely lack strict mechanisms to arrest the meiotic cell division. Contrary to animals and fungi that may arrest the cell cycle and abort meiosis once DSBs are not repaired, plants will progress through meiosis irrespective of such major defects [40].

(b) Meiosis and epigenetic reset

Meiosis and syngamy represent critical transitions between haploid and diploid phases in each generation. It has been suggested that a primary function of meiosis is to allow for epigenetic resetting in eukaryotes [99]. For instance, metazoan development is under the control of many epigenetic changes (cytosine methylation and chromatin marks) that are irreversibly maintained throughout life and must be reset twice each generation (at the n → 2n and 2nn transitions). This ensures proper development, the acquisition of parent-specific imprints, and may allow for mechanisms limiting the maximal number of possible successive mitoses (‘Hayflick limit’, reducing tumour development [99]). Some loci escape these resets, which can lead to transgenerational epigenetic inheritance [100]. This occurs much less frequently in animals than in plants (e.g. in Arabidopsis, demethylation is largely restricted to asymmetric CHH methylation sites, and contrary to mouse, does not occur on most symmetric CG and CHG methylation sites) [100]. Although the 2nn resetting occurs at or very close to meiosis in some cases (in female meiosis in animals), its timing may not be strictly tied to meiosis. For instance, it occurs pre-meiotically in the male germ line of animals (as shown in mice) or post-meiotically in male plant gametophytes (as shown in Arabidopsis) [100].

The evolutionary significance of these timing differences are poorly understood. Meiosis may simply not be the optimal time for epigenetic resetting. Many epigenetic pathways repress the activity of transposable elements (TEs), and so resetting epigenetic marks exposes the genome to mobilization of these elements, which may be particularly detrimental when producing gametes. In addition, meiosis may be specifically vulnerable to TE activity for several reasons [101,102]. These include (i) deficient synapsis and repair due to the reshuffling of the meiotic machinery towards TE-induced DSBs (ii) ectopic recombination among TEs, and (iii) interference with synapsis due to TE transcriptional activity. Alternative TE silencing mechanisms, such as those involving small RNAs, may have evolved to ensure proper TE control during epigenetic resetting. For example, these mechanisms involve piRNA and/or endo-siRNA in mammal male and female germlines, respectively [103], and transfer of siRNA from the central cell to the egg cell in plant female gametophytes [104]. It is also possible that stringent synapsis checkpoints evolved, in part, to prevent the formation of defective gametes due to TE activity, along with other possible causes of meiotic errors.

(c) Meiosis asymmetry

Symmetrical meiosis results in four viable gametes, whereas asymmetrical meiosis results in a single gamete. Symmetrical meiosis is ancestral and is found in male meiosis in animals, seed plants, ‘homosporous’ species (e.g. mosses, many ferns) and isogamous eukaryotes. Asymmetrical meiosis, on the other hand, has evolved multiple times, and occurs in female meiosis in animals, seed plants and some ciliates. The selective scenarios underlying the evolution of meiotic asymmetry are unresolved. In some cases, such as in ciliates, there is no requirement for four meiotic products, as sex occurs by the cytoplasmic exchange of haploid micronuclei (conjugation). In other cases, asymmetrical meiosis in females results in a large oocyte full of resources, which may favour the production of a single cell rather than four [66,105,106]. However, females could also achieve this symmetrically by undergoing fewer meioses. Therefore, is it possible that asymmetrical meiosis allows better control of resource allocation to oocytes, as symmetrical meiosis may not ensure an even distribution of resources across four meiocytes one difficulty here is that it is not clear why female control of resource allocation would be more efficient among meiocytes derived from the same or different meiosis. A solution may be that meiocytes must compete for resources during meiosis, so that a symmetrical female meiosis is vulnerable to SGEs that bias resource allocation in their favour, possibly by killing other products of meiosis [106]. Asymmetrical meiosis may therefore have evolved to suppress such costly competition within tetrads [107], but as discussed in the next section, it also opens the possibility of new conflicts [106]. Hence, the evolution of asymmetrical female meiosis is a question that remains not entirely resolved.

(d) Fairness of meiosis

A striking feature of meiosis is its apparent fairness: under Mendel's first Law of inheritance, each allele has a 50% chance of ending up in any given gamete. However, there are many SGEs that increase their chances above 50% by subverting the mechanism of meiosis. These SGEs fall into two classes. The first class is killer SGEs, which kill cells that have not inherited the element. In principle, such killers could operate during meiosis (the hypothetical ‘sister killers’ as discussed above), but the numerous killer SGEs that have been identified so far operate post-meiotically, e.g. by killing sibling sperm [108–111]. The second class consists of meiotic drivers that exploit the asymmetry of female meiosis discussed in the previous section. These elements achieve transmission in excess of 50% by preferentially moving into the meiotic products that will eventually become the eggs or megaspores [109,112]. There is a similarity between this kind of meiotic drive, where alleles preferentially go where resources are (i.e. the egg), and SGEs expressed later and biasing resource allocation in their favour [113]. Parents make decisions of allocations to offspring before the ‘meiotic veil of ignorance’, whereas offspring compete for resources ‘from behind the veil’ [114,115]. These genetic conflicts (between parent and offspring and between paternally and maternally derived alleles) are likely at the origin of parental imprints that differentially occur at male and female meiosis on some genes controlling embryo growth [114].

SGEs that undermine the fairness of meiosis provide explanations for otherwise puzzling observations. Perhaps most strikingly, centromere DNA regions often evolve rapidly, in contrast with what one would expect given their important and conserved function in meiosis. Henikoff et al. [116], therefore, proposed that expansion of repeat sequences in centromeric DNA produces a ‘stronger’ centromere, with increased kinetochore binding, which exhibits drive towards the future egg during meiosis I and, consequently, spreads in the population. Some of the best support for this hypothesis comes from a female meiotic driver in the monkeyflower Mimulus guttatus [117]. Although conclusive evidence for a direct centromere function of this element is lacking, it is physically associated with large centromere-specific satellite DNA arrays [118]. Female meiotic drive may also explain rapid karyotype evolution and the distribution of meta- versus acrocentric chromosomes [112] because Robertsonian fusion chromosomes (fusions of two acrocentric chromosomes into one metacentric) can behave like meiotic drivers and segregate preferentially into the future egg during meiosis I [119].

Other features of meiosis may be adaptations to suppress killer or meiotic drive SGEs. Such adaptations are expected, because these elements are generally costly for the rest of the genome (e.g. [108,120]). Defence against killer elements can be achieved by limiting gene expression. Accordingly, meiotic sex chromosome inactivation (MSCI, starting at pachytene of prophase I, figure 1) has been proposed to have evolved to control sex chromosome meiotic drive elements [121], and more generally this same principle may explain limited gene expression during meiosis and in its haploid products, as well as sharing of RNA and proteins among these cells. There is also evidence for rapid evolution and positive selection in the DNA-binding regions of centromere-associated proteins, which accords with the expectation of selection for countermeasures to limit preferential segregation of centromere drive elements towards the egg [106,116]. The evolution of holokinetic chromosomes may be an extreme form of defence against centromere drive [106].

5. Meiosis and recombination

A ubiquitous feature of meiosis is the exchange of genetic material between homologous chromosomes. While we have discussed arguments on its origin (see §2b), the maintenance of recombination is even more debated [122–124]. Here, we do not review this question, but discuss the evolutionary significance of patterns of recombination variation within and across species, as these present many mysteries connected to the functioning of meiosis.

(a) The number of crossovers per chromosome: constrained or not?

In many species, the number of COs per bivalent appears to follow highly constrained patterns, showing little variation compared to the variation of chromosome sizes, themselves spanning several orders of magnitude [125]. Within species, the correlation between genetic map length (in cM, with 50 cM being equivalent to 1 CO per bivalent) and physical length (in megabases, Mb) per chromosome is very strong (R 2 > 0.95) [126–131], and often has an intercept of approximately 50 cM, consistent with occurrence of one obligate CO per bivalent. There is direct evidence indicating that bivalents lacking a CO have an increased probability of non-disjunction, resulting in unviable or unfit aneuploid offspring [132,133]. Indeed, COs establish physical connections between homologues, promoting accurate disjunction by providing the tension needed for the bipolar spindle to establish [134–136]. Therefore, this constraint has likely led to the evolution of regulation of CO numbers per bivalent across the eukaryotes [137,138]. However, the reasons underlying the evolutionary persistence of this constraint are not well understood. In several species (e.g. Arabidopsis, [139]), the intercept is less than 50 cM, but the smallest chromosome is at least 50 cM, thus still consistent with one obligate CO. More decisively, many species are achiasmate (i.e. have an absence of recombination) in one sex [140], with alternative mechanisms to ensure proper disjunction of achiasmate bivalents [141,142]. This indicates that COs are not always obligatory and are maintained for reasons other than ensuring proper disjunction.

In addition to the obligate CO, additional CO events can occur within bivalents. The strong cM–Mb relationship within species indicates that the number of surplus COs correlates strongly with physical chromosome size (see above). However, the rate at which surplus COs are added per Mb (i.e. the slope of the correlation) varies strongly between species [125,131,143]. This may be partly explained by selection for different CO rates in different species [144–146]. The strong correlations observed within most species may be explained by variation in trans-acting factors, such as the locus RNF212 and its protein, which affects the propensity for DSBs to form surplus COs [147,148] indeed, the identification of loci affecting variation in CO rates indicates the potential for rapid evolution of CO rates within and between species [149].

A further constraint on bivalent disjunction may exist: the separation of different bivalents on the meiotic spindle may need to be collectively synchronized to avoid aneuploidy. If the number of COs correlates with the amount of tension exerted on the homologues, then a tight control of excess COs may minimize disjunction asynchrony. This hypothesis may explain the observation that some disjunction problems in humans occur in a global manner without involving effects driven by specific chromosomes [150–152]. Generally, high CO numbers are, on the other hand, not necessarily problematic with respect to proper disjunction [136,153].

(b) Crossover interference

A CO in one position may strongly reduce the likelihood of another CO occurring in the vicinity and/or on the same bivalent. This ‘crossover interference’ is widespread [125,136,154,155], but its function and mechanistic basis remains largely unknown. In many species, two classes of COs have been identified: Class I COs, which are sensitive to interference and Class II COs, which are not [156]. Class I COs are thought to play a major role in ensuring obligate COs, and so interference may limit the frequency or variance of COs, which may be important in ensuring proper disjunction [157]. For instance, as with autopolyploids (see above), increased interference may limit the number of COs to just one per chromosome, preventing aberrant multivalent segregation [54]. A variant of this idea is that interference is a mechanism to avoid COs occurring in close proximity, which might reduce cohesion between homologues [158] or slip and cancel each other out when they involve two or four non-interlocking chromatids, resulting in no CO occurring [159] however, these mechanisms do not explain long-distance interference. A further suggestion is that CO interference may be adaptive by breaking up genetic associations. First, adjacent COs may be avoided because they cancel their effects on genetic associations [160]. Second, it has been speculated that CO interference may reduce the chances of breaking up co-adapted gene complexes (supergenes) [157]. Some support for the idea that CO interference is not a purely mechanistic constraint comes from the fact that some species lack interference [154] and, more importantly, that there is some evidence suggesting that interference levels evolve in long-term evolution experiments in Drosophila [161].

(c) Differences in recombination rates between the sexes

In many species, CO rates and localization differ between male and female meioses, and these differences can vary in degree and direction even between closely related species [162–164]. The most extreme case is achiasmy, an absence of recombination in one sex, nearly always the heterogametic sex [162]. This may have evolved either as a side effect of selection to suppress recombination between the sex chromosomes [165,166], or as a way to promote tight linkage without suppressing recombination on the X or Z chromosomes [163]. More intriguing are the quantitative differences between males and females, known as heterochiasmy, which are found in many taxa, but whose mechanistic and evolutionary drivers are not yet fully understood. A number of explanations have been proposed, relating to mechanistic factors such as differences in chromatin structure [167–169], sexual dimorphism in the action of loci associated with CO rate (e.g. RNF212, [127,128,148]), and evolutionarily widespread processes such as sperm competition, sexual dimorphism and dispersal [162,170,171]. Some models point to a role of sex differences in selection during the haploid phase [172]. While a viable explanation in plants [163], there is little empirical support for this in animals [171], where meiosis in females is only completed after fertilization (i.e. there is no true haploid phase), and where only few genes are expressed in sperm. However, meiotic drive systems are often entirely distinct between males and females [173] and may be a primary cause of haploid selection [174]. These systems often require genetic associations between two loci (a distorter and responder, or a distorter and a centromere in males and females, respectively). These driving elements might thus be very important in shaping heterochiasmy patterns [107]. Indeed, COs in female meiosis are located closer to centromeres, which would be consistent with the view that this localization evolved to limit centromeric drive [175] (see also §4d). Similarly, meiotic drive in favour of recombinant chromatids has been detected in human female meiosis [67], which may limit centromere drive.

(d) The localization of crossovers and recombination hotspots

The localization of recombination events differs between species. In many species, recombination occurs in localized regions known as ‘recombination hotspots’ of approximately 1–2 kb in length [176–179], although some species (e.g. Caenorhabditis elegans and Drosophila) lack well-defined hotspots [180,181]. There are at least two types of hotspots (figure 3). The first type, probably ancestral, is found in fungi, plants, birds and some mammals these hotspots are temporally stable (up to millions of years) and concentrated near promoter regions and transcription start sites [178,182–185]. The second type is likely derived and is found in other mammals, including mice and humans, where the positioning of hotspots is determined by the zinc-finger protein PRDM9. This system differs in two respects from the former: first, it appears to direct DSBs away from regulatory regions [186], and second, mutations in the DNA-binding zinc-finger array change the sequence motif targeted by the protein, leading to rapid evolution of hotspot positions over short time scales [187,188]. This system is not present in all mammals: in dogs, hotspots target promoter regions [189], and the knock-out of Prdm9 in mouse makes recombination target promoter regions instead, underlining its derived nature [186].

Figure 3. Hypothetical genome sequence containing three genes showing the distribution of ancient recombination hotspots in most model species (a) compared with derived PRDM9-mediated recombination hotspots (b). Studies in fungi, plants, birds and dogs indicate that ancestral hotspots are stable over long evolutionary timescales (up to millions of years) and concentrate at promoter regions and transcription start sites (and at stop sites in some species). These start and stop sites for each gene are indicated in yellow and red blocks, with their introns and exons represented by lines and black blocks, respectively. PRDM9-mediated hotspots are found in some mammals, including humans and mice, and are directed away from promotor regions. The DNA-binding zinc-finger in the PRDM9 protein targets specific sequence motifs mutations in the zinc-finger array change the targeted motif, leading to rapid evolution of hotspot positions and an absence of hotspot conservation over short evolutionary timescales (at the population and species level).

The evolutionary significance of both kinds of hotspots remains unclear. For the first type, the positions of hotspots may be caused by chromatin accessibility in transcribed regions or have evolved to favour recombination in gene rich regions (where it might be worth reducing genetic association). However, this does not clearly account for their precise location in regulatory regions. Another possibility might be that the co-occurrence of both COs and gene-conversion events (i.e. where resolution of DSBs without CO is achieved by exchanging small segments of DNA) specifically in regulatory regions could repress enhancer runaway, a mechanism that can lead to suboptimal expression levels [190]. The evolutionary significance of the second kind of hotspot is similarly elusive. These hotspots are self-destructing because the target sequence motifs are eroded by biased gene conversion (BGC) during DSB repair [191]. This leads to a ‘hotspot paradox’: how can hotspots and recombination be maintained in the long term in the face of BGC [192]? A possible solution is that trans-acting factors like PRDM9 may mutate sufficiently fast to constantly ‘chase’ new and frequent targets (hotspots), switching to new ones when these targets become rare due to BGC [193]. This ‘Red Queen’ model does not require strong stabilizing selection on the number of COs, and closely mimics the pattern of hotspot turnover observed in some cases [194]. However, this model does not explain how the second kind of hotspots evolved in the first place, as when it arose proper segregation was presumably already ensured by the first kind of hotspots (which, as seen in mice, are still active). Also, it does not explain why PRDM9 action is self-destructing: there is no necessity to induce DSBs exactly at the position of the target sequence for a trans-acting factor. In fact, there is no logical necessity to rely on a target sequence to maintain one CO per chromosome, as fixed chromosomal features could serve this purpose. It is worth noting here that recruiting promoter sequences for this purpose (as found for hotspots of the first kind) would be very efficient, as these sequences are highly stable and dispersed in the genome on all chromosomes. There is also no evidence so far that targeted binding motifs of PRDM9 correspond to some SGEs whose elimination would be beneficial. Overall, while spectacular progress has been made recently in elucidating hotspot mechanisms in detail (and patterns in recombination landscapes), there are still major gaps in our understanding of their evolutionary significance.

6. Conclusion

The evolutionary significance of meiosis has often been interpreted in an oversimplified manner, restricted mainly to the direct (DSB repair, proper disjunction) or indirect (genetic associations) effects of meiotic recombination. Yet, many features of meiosis are unlikely to be explained by effects of recombination alone, and the fields of cellular and molecular biology are uncovering new meiotic features at a high rate. One of the main take-home messages of this review is that many, if not most features of meiosis are still awaiting an evolutionary explanation. Nonetheless, the recent advances in all detailed aspects of meiosis now offer the chance to investigate these questions in a far more comprehensive manner. This will require continued dialogue between cell, molecular and evolutionary biologists (as advocated e.g. in [195]), and perhaps also the realization that similarities between features may in fact have different evolutionary explanations (e.g. different kinds of hotspots).

One of the most salient themes in most meiosis mysteries is the impact of genetic conflicts and SGEs. As for the evolution of genome size and structure, their impact is probably central [196], but in many cases they remain hypothetical and difficult to demonstrate and study directly: many SGEs reach fixation quickly and leave almost no visible footprint. Showing that some meiotic features evolved to control SGEs represents an even greater challenge. Indeed, if successful, such features would prevent these SGEs from spreading, further limiting their detection. In addition, demonstrating a role in SGEs control requires ruling out that these features evolved for more mechanistic and simpler alternatives. This is usually extremely difficult, as many ad hoc mechanistic constraints can be imagined.


Asexual Microorganisms and Animals

A wide variety of microorganisms reproduce asexually. Protozoans, bacteria and a group of algae called diatoms reproduce through fission. The simple microscopic animals known as cnidaria, and the annelids, also called ringworms, reproduce through fragmentation. Biologists have discovered nearly 70 species of vertebrates that can reproduce parthogenetically, including frogs, chickens, turkeys, Komodo dragons and hammerhead sharks.


NEET DPP Biology Ch-23 Reproduction in Organism

Answer (b) Cloning is a technique by which genetically same individuals can be produced without including any sexual reproduction eg. Dolly sheep.

Q.3. Natural parthenogenesis occurs in:
(a) Protozoans
(b) Earthworm
(c) All insects
(d) Honeybee

Q.4. Retention of larval characters even after sexual maturity is called
(a) Parthenogenesis
(b) Ontogenesis
(c) Paedogenesis
(d) Neoteny

Q.5. Asexual reproduction is an effective strategy in stable environments because
(a) gametogenesis is most efficient under these conditions.
(b) the offspring, genetically identical to their parents, are preadapted to the environment.
(c) asexual parthenogenesis produces a large amount of genetic diversity.
(d) animal cells tend to be more totipotent under stable conditions.

Answer (b) The parents that have survived to reproduce asexually are able to survive in the current stable environment. Therefore, the offspring should be preadapted for this stable environment.

Q.6. If you compared the genetic makeup of an animal produced by parthenogenesis with that of its mother, which of the following would you expect?
(a) About 100 percent genetic similarity
(b) About 50 percent genetic similarity
(c) No genetic similarity
(d) Parthenogenetic animals have no mother

Answer (a) Species that exhibit parthenogenesis develop from unfertilized eggs produced by the mother. Therefore, the genetic make-up should be 100 percent the same as the mother. environment.

Q.7. Which of the following statements about animals that utilize external fertilization is false?
(a) They are divided equally between terrestrial and aquatic species.
(b) Many produce large numbers of gametes to ensure successful reproduction.
(c) The behaviours associated with mating are often highly synchronized.
(d) The probability of any one egg being fertilized and developing into an adult can be low.

Answer (a) Since external fertilization can only take place in an aquatic habitat, there are no terrestrial animals that use it.

Q.8. Which of the following statements about animal reproduction is false ?
(a) Species that reproduce sexually cannot also reproduce asexually.
(b) Viviparity, but not ovoviviparity, is common in mammals.
(c) Male insects can remove spermatophores deposited in a female by other males.
(d) Oogenesis and spermatogenesis both occur in simultaneous hermaphrodites.

Answer (a) Many animals reproduce both by asexual and sexual means.

Q.9. Which of the following animals qualifies as a sexually reproducing, oviparous species ?
(a) Human
(b) Chicken
(c) Kangaroo
(d) Sea star

Answer (b) All these animals can reproduce sexually. However, only the chicken lays an external egg.

Q.10. Benefits of asexual reproduction include all of the following except
(a) it often allows for the production of many more offspring at the same time
(b) it is advantageous in changing environments in which population variety is the key to successful propagation of a species
(c) it is easier in certain environments to have offspring without searching for a mate
(d) allowing the conservation of resources otherwise allocated to finding mates and performing ritualized courtship.

Answer (b) All of these statements concerning asexual reproduction are correct, except that asexual reproduction is best in favorable, stable environments, ones that don’t change rapidly. The reason for this is that asexual reproduction, in contrast to its sexual counterpart, results in the formation of identical offspring. Although asexual organisms can often produce many more offspring in a single reproductive event than sexual organisms, these asexually produced young do not usually have the genetic variation caused by meiosis and crossingover to be able to survive a rapidly changing environment or times of environmental stress.

Q.11. All the ‘eyes’ of a potato tuber are taken out and it is sown in the ground normally. New plants will
(a) Not emerge
(b) Emerge normally
(c) Be weaker
(d) Be healthier

Answer (a) Buds in ‘eyes’ form new plants.

Q.12. Basal half of an onion bulb is removed and upper half is sown in the ground. New plant will
(a) Emerge normally
(b) Not emerge
(c) Be without leaves
(d) Be without flowers

Answer (b) Bud giving rise to new plant is present towards base.

Q.13. A small portion of cane-sugar stem between the two successive nodes is cut off and sown in the soil normally. New plants will
(a) Be formed normally
(b) Not be formed
(c) Be without juice
(d) Without nodes

Answer (b) New plants in cane-sugar are formed from nodes which are absent.

Q.14. A cutting of rose plant is thoroughly waxed and planted in the soil normally, It will form
(a) New rose plant
(b) A dead piece of rose stem after some time
(c) A rose plant of improved variety
(d) None of these

Answer (b) Water absorption & gaseous exchange stop due to presence of wax

Q.15. When an ovary develops into a fruit, without fertilization, it is called
(a) apospory
(b) apogamy
(c) parthenocarpy
(d) porogamy

Answer (c) Parthenocarpy is the development of a fruit without the formation of seeds as a result of lack of pollination, lack of fertilization and lack of development. This condition can be artificially induced by application of hormones.

Q.16. Asexual reproduction is the best strategy for plants
(a) that are well adapted to their stable environment.
(b) as winter approaches
(c) when new genes must be introduced
(d) that have underground stems.

Q.17. Bamboo reproduces by
(a) rhizomes
(b) tubers
(c) corms
(d) stolons

Q.18. Grafting is an example of asexual reproduction. Which of the following choices is an example of asexual reproduction involving nonvegetative parts of a plant ?
(a) Apomixis
(b) Production of corms
(c) Production of bulbs
(d) Production of rhizomes

Q.19. What is necessary for successful grafting to occur ?
(a) Each section must be able to form roots.
(b) The grafted section must be able to form seeds.
(c) Fusion of the two vascular tissues must occur.
(d) Fusion of the two cambial tissues must occur.

Q.20. Banana fruits are seedless, because
(a) auxins are sprayed for rapid development of fruit.
(b) vegetative propagation of plant.
(c) they are triploid plants.
(d) fruits are artificially ripened.

Answer (c) Most of banana varieties are triploid and triploidy is associated with seedlessness.

Q.21. Consider the following statements and choose the correct option.
(i) The genetic constitution of a plant is unaffected in vegetative propagation.
(ii) Rhizome in ginger serves as an organ of vegetative reproduction.
(iii) Totipotency of cells enables us to micropropagate plants.
(a) Statements (i) and (ii) alone are true
(a) Statements (ii) and (iii) alone are true
(c) Statement (ii) alone is true
(d) All the three statements (i), (ii) and (iii) are true

Q.22. Plants identical to mother plants can be obtained from
(a) seeds
(b) stem cutting
(c) Both (a) and (b)
(d) None of these

Q.23. Ploidy of ovary, anther, egg, pollen, male gamete and zygote are respectively-
(a) 2n, 2n, n, 2n, n, 2n
(b) 2n, 2n, n, n, n, 2n
(c) 2n, n, n, n, n, n
(d) 2n, 2n, n, 2n, 2n, 2n

Q.24. Offsprings of oviparous animals are at greater risk as compared to offsprings of viviparous animals because-
(a) Proper embryonic care and protection is lesser
(b) Embryo is not developed
(c) Progenies are with more variation
(d) Progenies are larger

Q.25. The parameter(s) of senescence or old age is/are-
(a) End of the reproductive phase
(b) Concomitant change in body (like slowing metabolism)
(c) Failure of metabolism decreases
(d) Both (a) and (b)

Q.26. The terms homothallic and monoecious are used to denote
(a) bisexual condition
(b) unisexual condition
(c) staminate flowers
(d) pistillate flowers

Answer (a) Homothallic and monoecious are terms used to denote bisexual condition. The example indudes fungi and plants. Heterothallic and dioecious are terms used to denote unisexual condition.

Q.27. During regeneration, modification of an organ to other organ is known as
(a) Morphogenesis
(b) Epimorphosis
(c) Morphallaxis
(d) Accretionary growth

Answer (b) Morphallaxis is a mechanism of regeneration involving reorganization of body cells. In epimorphosis, new cells proliferate from the surface of the wound to form the missing structure. In accretionary growth some specialized cells retain the ability to divide and produce new cells to replace the worn-out.

Q.28. Cells become variable in morphology and function in different regions of the embryo. The process is
(a) differentiation
(b) metamorphosis
(c) organisation
(d) rearrangement

Answer (a) Cells become variable in shape, size & getting their specialization for the formation of particular tissue or organ in future foetus. They place themselves at some specific regions in embryo for further organogeny.

Q.29. Earthworms, sponges, tapeworms are
(a) bisexual animals
(b) unisexual animals
(c) hermaphrodites
(d) Both (a) and (c)

Answer (d) Earthworm, sponges, tapeworms are bisexual animals and hermaphrodites as they prossess both male and female reproductive organs.

Q.30. The site of origin of the new plantlets in potato, dahlia, ginger and banana is-
(a) Floral buds present on stem
(b) Internodes of modified stem
(c) Nodes of modified stem
(d) Adventitious buds present on root

Q.31. Among the following which one is not a method of vegetative propagation?
(a) Budding
(b) Layering
(c) Sowing
(d) Tissue culture

Answer (c) Sowing is related with sexual reproduction.

Q.32. Vegetative propagation in mint occurs by:
(a) offset
(b) rhizome
(c) sucker
(d) runner

Answer (c) Vegetative propagation in mint occurs through sucker. Vegetative reproduction is a type of asexual reproduction for plants, and is also called vegetative propagation, vegetative multiplication, or vegetative cloning. It is a process by which new plant “individuals” arise or are obtained without production of seeds or spores. It is a natural process in many plant species (as well as non-plant organisms such as bacteria and fungi) and used or encouraged by horticulturists to obtain quantities of economically valuable plants. A related technique used in cultivation is tissue culture, which involves vegetative reproduction under sterile conditions.

Q.33. What is common between vegetative reproduction and apomixis?
(a) Both are applicable to only dicot plants
(b) Both bypass the flowering phase
(c) Both occur round the year
(d) Both produces progeny identical to the parent

Answer (d) Vegetative reproduction and apomixis, both are asexual methods of reproduction, which gives the progeny genetically similar to parent.

Q.34. Individuals of a clone-
(a) Are genetically similar but morphologically different
(b) Are morphologically similar but genetically different
(c) Are morphologically and genetically similar
(d) Are genetically and phenotypically different

Q.35. Some organisms are capable of asexual or sexual reproduction. Under favourable conditions, reproduction proceeds asexually. When conditions become more stressful reproduction switchess to a sexual mode. Why?
(a) Sexual reproduction is simple and more rapid allowing larger numbers of offspring to be produced
(b) Sexual reproduction requires two separate individuals, who can mutually provide nutrient support during stress
(c) Sexual reproduction produces individuals with new combinations of recombined chromosomes increasing diversity
(d) Asexual reproduction requires more energy

Q.36. Apomixis in plant means development of a plant
(a) from root cuttings
(b) without fusion of gametes
(c) from fusion of gametes
(d) from cuttings of stem

Q.37. Which of the following is not vegetative propagule ?
(a) Rhizome and sucker
(b) Tuber and offset
(c) Bulbil (e.g. in Agave), leaf buds and bulb
(d) Antherozoid

Q.38. Which of the following is false about external fertilization?
(a) Organisms showing external fertilization exhibit great synchrony between the sexes and release gametes.
(b) Gametes are produced in large number in water to enhance the chances of fertilization.
(c) A large number of gametes are wasted.
(d) A major advantage is that the offspring are protected from predators and there is a great chance of their survival upto adulthood.

Q.39. Modified stem present in Gladiolus is:
(a) bulb
(b) rhizome
(c) corm
(d) bulbil

Q.40. Which of the following are seasonal breeders?
(a) Frogs
(b) Birds
(c) Lizards
(d) All of these

Q.41. Select the wrong statement:
(a) Anisogametes differ either in structure, function or behaviour.
(b) In oomycetes female gamete is smaller and motile, while male gamete is larger and non-motile.
(c) Chalmydomonas exhibits both isogamy and anisogamy and Fucus shows oogamy.
(d) Isogametes are similar in structure, function and behaviour.

Answer (b) In oomycetes female gamete is large and non motile while male gamete is small & motile.

Q.42. Monoecious plant of Chara shows occurrence of :
(a) stamen and carpel of the same plant
(b) upper antheridium and lower oogonium on the same plant
(c) upper oogonium and lower antheridium on the same plant
(d) antheridiophore and archegoniophore on the same plant

Answer (c) Male sex organ is called antheridium or globule while female sex organ is called oogonium. They develop on the same branchlet in the same plant in chara.

Q.43. Which of the following statement(s) is/are false about internal fertilization?
(i) Male gametes are motile.
(ii) Male gametes are non-motile.
(iii) Male gametes are produced in large number.
(iv) Male gametes are produced in small number.
(v) There is a significant reduction in the number of eggs produced.
(a) (i), (iii) and (v)
(b) (iii) and (iv)
(c) (ii) and (iv)
(d) Only (v)

Q.44. Syngamy can occur outside the body of the organism in
(a) Fungi
(b) Mosses
(c) Algae
(d) Ferns

Answer (c) In most aquatic organisms, such as a majority of algae and fishes as well as amphibians, syngamy occurs in the external medium (water), i.e., outside the body of the organism. This type of gametic fusion is called external fertilisation.

Q.45. Select the correct sequence from the following.
(i) Juvenile phase → Senescent phase → Reproductive phase
(ii) Juvenile phase → Reproductive phase → Senescent phase
(iii) Reproductive phase → Juvenile phase → Senescent phase
(iv) Vegetative phase → Reproductive phase → Senescent phase
(a) (i) and (ii)
(b) (i) and (iv)
(c) (iii) and (iv)
(d) (ii) and (iv)