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Difference In Telomeres Between A Thale Cress Plant And A Methuselah Tree

Difference In Telomeres Between A Thale Cress Plant And A Methuselah Tree



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From what I have read and understood telomeres cap off how many times a cell can divide before it can no longer divide and that is what causes aging.

A thale cress plant apparently has a life cycle of 6 weeks before it dies, while the Methuselah tree has set the record at over 4,800+ years.

  • What is the difference between these two plants regarding their telomeres?
  • Is the cell division rate different between both plants instead of a telomeres difference?
  • If given ideal conditions what could the potential life-span of a Methuselah tree be?

Btw you don't have to answer all these. Just curious and looking for some insight into why the stark difference between both plants' aging.


Telomeres do not "cause" ageing as such - although you are right that they limit the number of times a somatic cell can divide.

Each time a cell divides the chromosomes are replicated in an imperfect way, and as such a small amount of DNA is lost from the end of the chromosome during each round of cell division. Telomeres are just extensions to the chromosomes, so that the DNA that is lost is not important. When a cell runs out of telomeres it ceases to divide.

However it is a big generalization to say that telomere length is therefore correlated with lifespan. For instance a recent study has shown that mice have telomeres 5-10x the length of humans, but live 30x shorter (1).

Inter-species differences in lifespan

Whilst I am no expert on these 2 plants, it is quite an obvious thing to say that organisms age at different rates. Mice, for instance, live 3 years in a protective environment, whereas humans can live 30x that. The difference is down to the life strategies acquired by natural selection: it is more advantageous for a mouse to develop very fast to reproductive maturity and have as many offspring as possible, because there is a very high mortality rate for mice. Conversely for humans, in takes more than a decade for us to reach reproductive maturity! Our "rate" of development/ageing is lower, which is completely related to our reproductive strategies.

I imagine that a very similar story is true for plants - for some it will be advantageous for them to "live fast, die young", whereas for others the best strategy might be to take a long time creating less, but more robust, offspring.

At the molecular level

I have answered a related question previously (Do trees age on a microscopic level?) which might be of interest to you.

In short, the phenotype of ageing is associated with irreparable damage at the molecular level - be it to DNA, organelles, protein aggregations… you name it! Organisms that have a higher metabolic rate, such as mice, accumulate this damage faster. This gives them the benefit of being able to reproduce much earlier, at the cost of reduced overall lifespan.

Plants age slightly differently, but at the genetic level much the same repair pathways are conserved throughout the kingdoms of life, with some alterations of course - for instance plants seem almost completely immune to cancer (2)!

As I argue in the linked to question above, plants have a potentially indefinite replicative potential, as every cell has the capability of regenerating its telomeres, because plants do not have a conserved germ line as animals do - any cutting from a plant has the potential to create an entirely new plant. This varies from plant to plant of course, and plants are still susceptible to damage, they just have modified/adapted repair pathways to cope with their specific lifestyle.

You ask whether it is the telomeres or the "rate" that differs between the plants - the answer is almost certainly a combination of both, amongst the other myriad of factors that influence lifespan!

Refs

  1. Colado, et al, 2013. Telomere dynamics in mice and humans. Seminars in hematology [PubMed]
  2. Doonan & Sablowski, 2010. Walls around tumours - why plants do not develop cancer. Nature reviews cancer [DOI]

Understanding Brassicaceae evolution through ancestral genome reconstruction

Brassicaceae is a family of green plants of high scientific and economic interest, including thale cress (Arabidopsis thaliana), cruciferous vegetables (cabbages) and rapeseed.

Results

We reconstruct an evolutionary framework of Brassicaceae composed of high-resolution ancestral karyotypes using the genomes of modern A. thaliana, Arabidopsis lyrata, Capsella rubella, Brassica rapa and Thellungiella parvula. The ancestral Brassicaceae karyotype (Brassicaceae lineages I and II) is composed of eight protochromosomes and 20,037 ordered and oriented protogenes. After speciation, it evolved into the ancestral Camelineae karyotype (eight protochromosomes and 22,085 ordered protogenes) and the proto-Calepineae karyotype (seven protochromosomes and 21,035 ordered protogenes) genomes.

Conclusions

The three inferred ancestral karyotype genomes are shown here to be powerful tools to unravel the reticulated evolutionary history of extant Brassicaceae genomes regarding the fate of ancestral genes and genomic compartments, particularly centromeres and evolutionary breakpoints. This new resource should accelerate research in comparative genomics and translational research by facilitating the transfer of genomic information from model systems to species of agronomic interest.


Features of chromosome 2

Chromosome 2 is acrocentric and was originally estimated to be 13.5 Mb in length from the YAC-based physical map 12 . Sequencing was initiated using bacterial artificial chromosomes (BACs) that had been placed onto the physical map of chromosome 2 by hybridization to YACs and genetic markers. Subsequently, BAC end sequences and BAC fingerprint data 13 allowed extension from these initial seed points and completion of the entire chromosome. A total of 257 BAC and P1 clones (including 5 BACs completed by other groups) were sequenced to produce over 24 Mb of finished sequence, which has been assembled as two contigs terminating in blocks of 180-bp repeats. These repeats represent the inner boundaries of our finished sequence.

The upper (short) arm, measured from the lower end of the NOR to the centromeric 180 bp repeats, is 3.6 Mb. The northern-most BAC (F23H14) in this contig contains a single rDNA repeat unit that is oriented in the same direction as the telomere-proximal repeat, suggesting that all the rDNA units are arranged in a head-to-tail fashion running 5′ to 3′ from the telomere towards the centromere 14 . Immediately adjoining the rDNA is about 60 kb of highly repetitive sequence containing many transposons. The lower (long) arm from centromere to telomere is 16 Mb. The sequence of the southern-most BAC in this contig (F11L15) is joined by a small PCR fragment to the A. thaliana sequence in pAtT51, a telomere-containing clone previously mapped to TEL2S 15 (http://genome-www.stanford.edu/Arabidopsis/ww/Nov98RImaps/index.html). Within this region, a putative transcriptional co-activator gene was identified. Because pAtT51 was derived from the Landsberg ecotype, the last 2.5 kb of this contig may not be identical to the sequence in the Columbia ecotype.

At 19.6 Mb, the overall length of chromosome 2 (excluding the NOR and centromere) is 45% larger than the original estimate. This difference is due in part to gaps in the YAC-based physical map and to a deletion in one of the YACs (see Fig. 1). Similar increases in the actual physical lengths of the other A. thaliana chromosomes indicate that the genome size is actually 130–140 Mb, rather than the earlier estimates of 70–100 Mb.

Using sequence information for 37 markers from the Arabidopsis Recombinant Inbred genetic map (http://genome-www.stanford.edu/Arabidopsis/ww/Nov98RImaps/index.html), the relationship between physical and genetic distances along the chromosome was examined. As the centromere represents a discontinuity, regression analysis for the two arms was performed separately. The ratios of physical to genetic distance on the short arm and long arm were not significantly different, with values of 244 kb cM -1 and 223 kb cM -1 , respectively (Fig. 2a). Although these two values indicate a similar overall rate of recombination along the two arms of the chromosome, local distortions are evident which may reflect chromosomal rearrangements between the two ecotypes used to construct the mapping population (Fig. 2b).

a, Physical distance (in kb) is plotted against position of each marker on the Recombinant Inbred (RI) map. Data above and below the centromere are treated as separate sets for regression analysis. Note that genetic distance is measured from the telomere (TEL2N), whereas physical distance is measured from the bottom of the NOR. b, The ratio of physical to genetic distance between successive pairs of markers is plotted against map position for each marker pair on the RI map.


Contents

Types Edit

Polyploid types are labeled according to the number of chromosome sets in the nucleus. The letter x is used to represent the number of chromosomes in a single set:

  • haploid (one set 1x)
  • diploid (two sets 2x)
  • triploid (three sets 3x), for example sterile saffron crocus, or seedless watermelons, also common in the phylumTardigrada[5]
  • tetraploid (four sets 4x), for example Salmonidae fish, [6] the cotton Gossypium hirsutum[7]
  • pentaploid (five sets 5x), for example Kenai Birch (Betula papyrifera var. kenaica)
  • hexaploid (six sets 6x), for example wheat, kiwifruit[8]
  • heptaploid or septaploid (seven sets 7x)
  • octaploid or octoploid, (eight sets 8x), for example Acipenser (genus of sturgeon fish), dahlias
  • decaploid (ten sets 10x), for example certain strawberries
  • dodecaploid (twelve sets 12x), for example the plants Celosia argentea and Spartina anglica[9] or the amphibian Xenopus ruwenzoriensis.

Classification Edit

Autopolyploidy Edit

Autopolyploids are polyploids with multiple chromosome sets derived from a single taxon.

Two examples of natural autopolyploids are the piggyback plant, Tolmiea menzisii [10] and the white sturgeon, Acipenser transmontanum. [11] Most instances of autopolyploidy result from the fusion of unreduced (2n) gametes, which results in either triploid (n + 2n = 3n) or tetraploid (2n + 2n = 4n) offspring. [12] Triploid offspring are typically sterile (as in the phenomenon of 'triploid block'), but in some cases they may produce high proportions of unreduced gametes and thus aid the formation of tetraploids. This pathway to tetraploidy is referred to as the “triploid bridge”. [12] Triploids may also persist through asexual reproduction. In fact, stable autotriploidy in plants is often associated with apomictic mating systems. [13] In agricultural systems, autotriploidy can result in seedlessness, as in watermelons and bananas. [14] Triploidy is also utilized in salmon and trout farming to induce sterility. [15] [16]

Rarely, autopolyploids arise from spontaneous, somatic genome doubling, which has been observed in apple (Malus domesticus) bud sports. [17] This is also the most common pathway of artificially induced polyploidy, where methods such as protoplast fusion or treatment with colchicine, oryzalin or mitotic inhibitors are used to disrupt normal mitotic division, which results in the production of polyploid cells. This process can be useful in plant breeding, especially when attempting to introgress germplasm across ploidal levels. [18]

Autopolyploids possess at least three homologous chromosome sets, which can lead to high rates of multivalent pairing during meiosis (particularly in recently formed autopolyploids, also known as neopolyploids) and an associated decrease in fertility due to the production of aneuploid gametes. [19] Natural or artificial selection for fertility can quickly stabilize meiosis in autopolyploids by restoring bivalent pairing during meiosis, but the high degree of homology among duplicated chromosomes causes autopolyploids to display polysomic inheritance. [20] This trait is often used as a diagnostic criterion to distinguish autopolyploids from allopolyploids, which commonly display disomic inheritance after they progress past the neopolyploid stage. [21] While most polyploid species are unambiguously characterized as either autopolyploid or allopolyploid, these categories represent the ends of a spectrum of divergence between parental subgenomes. Polyploids that fall between these two extremes, which are often referred to as segmental allopolyploids, may display intermediate levels of polysomic inheritance that vary by locus. [22] [23]

About half of all polyploids are thought to be the result of autopolyploidy, [24] [25] although many factors make this proportion hard to estimate. [26]

Allopolyploidy Edit

Allopolyploids or amphipolyploids or heteropolyploids are polyploids with chromosomes derived from two or more diverged taxa.

As in autopolyploidy, this primarily occurs through the fusion of unreduced (2n) gametes, which can take place before or after hybridization. In the former case, unreduced gametes from each diploid taxa – or reduced gametes from two autotetraploid taxa – combine to form allopolyploid offspring. In the latter case, one or more diploid F1 hybrids produce unreduced gametes that fuse to form allopolyploid progeny. [27] Hybridization followed by genome duplication may be a more common path to allopolyploidy because F1 hybrids between taxa often have relatively high rates of unreduced gamete formation – divergence between the genomes of the two taxa result in abnormal pairing between homoeologous chromosomes or nondisjunction during meiosis. [27] In this case, allopolyploidy can actually restore normal, bivalent meiotic pairing by providing each homoeologous chromosome with its own homologue. If divergence between homoeologous chromosomes is even across the two subgenomes, this can theoretically result in rapid restoration of bivalent pairing and disomic inheritance following allopolyploidization. However multivalent pairing is common in many recently formed allopolyploids, so it is likely that the majority of meiotic stabilization occurs gradually through selection. [19] [21]

Because pairing between homoeologous chromosomes is rare in established allopolyploids, they may benefit from fixed heterozygosity of homoeologous alleles. [28] In certain cases, such heterozygosity can have beneficial heterotic effects, either in terms of fitness in natural contexts or desirable traits in agricultural contexts. This could partially explain the prevalence of allopolyploidy among crop species. Both bread wheat and Triticale are examples of an allopolyploids with six chromosome sets. Cotton, peanut, or quinoa are allotetraploids with multiple origins. In Brassicaceous crops, the Triangle of U describes the relationships between the three common diploid Brassicas (B. oleracea, B. rapa, and B. nigra) and three allotetraploids (B. napus, B. juncea, and B. carinata) derived from hybridization among the diploid species. A similar relationship exists between three diploid species of Tragopogon (T. dubius, T. pratensis, and T. porrifolius) and two allotetraploid species (T. mirus and T. miscellus). [29] Complex patterns of allopolyploid evolution have also been observed in animals, as in the frog genus Xenopus. [30]

Aneuploid Edit

Organisms in which a particular chromosome, or chromosome segment, is under- or over-represented are said to be aneuploid (from the Greek words meaning "not", "good", and "fold"). Aneuploidy refers to a numerical change in part of the chromosome set, whereas polyploidy refers to a numerical change in the whole set of chromosomes. [31]

Endopolyploidy Edit

Polyploidy occurs in some tissues of animals that are otherwise diploid, such as human muscle tissues. [32] This is known as endopolyploidy. Species whose cells do not have nuclei, that is, prokaryotes, may be polyploid, as seen in the large bacterium Epulopiscium fishelsoni. [33] Hence ploidy is defined with respect to a cell.

Monoploid Edit

A monoploid has only one set of chromosomes and the term is usually only applied to cells or organisms that are normally diploid. The more general term for such organisms is haploid.

Temporal terms Edit

Neopolyploidy Edit

A polyploid that is newly formed.

Mesopolyploidy Edit

That has become polyploid in more recent history it is not as new as a neopolyploid and not as old as a paleopolyploid. It is a middle aged polyploid. Often this refers to whole genome duplication followed by intermediate levels of diploidization.

Paleopolyploidy Edit

Ancient genome duplications probably occurred in the evolutionary history of all life. Duplication events that occurred long ago in the history of various evolutionary lineages can be difficult to detect because of subsequent diploidization (such that a polyploid starts to behave cytogenetically as a diploid over time) as mutations and gene translations gradually make one copy of each chromosome unlike the other copy. Over time, it is also common for duplicated copies of genes to accumulate mutations and become inactive pseudogenes. [34]

In many cases, these events can be inferred only through comparing sequenced genomes. Examples of unexpected but recently confirmed ancient genome duplications include baker's yeast (Saccharomyces cerevisiae), mustard weed/thale cress (Arabidopsis thaliana), rice (Oryza sativa), and an early evolutionary ancestor of the vertebrates (which includes the human lineage) and another near the origin of the teleost fishes. [35] Angiosperms (flowering plants) have paleopolyploidy in their ancestry. All eukaryotes probably have experienced a polyploidy event at some point in their evolutionary history.

Other similar terms Edit

Karyotype Edit

A karyotype is the characteristic chromosome complement of a eukaryote species. [36] [37] The preparation and study of karyotypes is part of cytology and, more specifically, cytogenetics.

Although the replication and transcription of DNA is highly standardized in eukaryotes, the same cannot be said for their karyotypes, which are highly variable between species in chromosome number and in detailed organization despite being constructed out of the same macromolecules. In some cases, there is even significant variation within species. This variation provides the basis for a range of studies in what might be called evolutionary cytology.

Homoeologous chromosomes Edit

Homoeologous chromosomes are those brought together following inter-species hybridization and allopolyploidization, and whose relationship was completely homologous in an ancestral species. For example, durum wheat is the result of the inter-species hybridization of two diploid grass species Triticum urartu and Aegilops speltoides. Both diploid ancestors had two sets of 7 chromosomes, which were similar in terms of size and genes contained on them. Durum wheat contains a hybrid genome with two sets of chromosomes derived from Triticum urartu and two sets of chromosomes derived from Aegilops speltoides. Each chromosome pair derived from the Triticum urartu parent is homoeologous to the opposite chromosome pair derived from the Aegilops speltoides parent, though each chromosome pair unto itself is homologous.

Animals Edit

Examples in animals are more common in non-vertebrates [38] such as flatworms, leeches, and brine shrimp. Within vertebrates, examples of stable polyploidy include the salmonids and many cyprinids (i.e. carp). [39] Some fish have as many as 400 chromosomes. [39] Polyploidy also occurs commonly in amphibians for example the biomedically-important genus Xenopus contains many different species with as many as 12 sets of chromosomes (dodecaploid). [40] Polyploid lizards are also quite common, but are sterile and must reproduce by parthenogenesis. [ citation needed ] Polyploid mole salamanders (mostly triploids) are all female and reproduce by kleptogenesis, [41] "stealing" spermatophores from diploid males of related species to trigger egg development but not incorporating the males' DNA into the offspring. While mammalian liver cells are polyploid, rare instances of polyploid mammals are known, but most often result in prenatal death.

An octodontid rodent of Argentina's harsh desert regions, known as the plains viscacha rat (Tympanoctomys barrerae) has been reported as an exception to this 'rule'. [42] However, careful analysis using chromosome paints shows that there are only two copies of each chromosome in T. barrerae, not the four expected if it were truly a tetraploid. [43] This rodent is not a rat, but kin to guinea pigs and chinchillas. Its "new" diploid (2n) number is 102 and so its cells are roughly twice normal size. Its closest living relation is Octomys mimax, the Andean Viscacha-Rat of the same family, whose 2n = 56. It was therefore surmised that an Octomys-like ancestor produced tetraploid (i.e., 2n = 4x = 112) offspring that were, by virtue of their doubled chromosomes, reproductively isolated from their parents.

Polyploidy was induced in fish by Har Swarup (1956) using a cold-shock treatment of the eggs close to the time of fertilization, which produced triploid embryos that successfully matured. [44] [45] Cold or heat shock has also been shown to result in unreduced amphibian gametes, though this occurs more commonly in eggs than in sperm. [46] John Gurdon (1958) transplanted intact nuclei from somatic cells to produce diploid eggs in the frog, Xenopus (an extension of the work of Briggs and King in 1952) that were able to develop to the tadpole stage. [47] The British scientist J. B. S. Haldane hailed the work for its potential medical applications and, in describing the results, became one of the first to use the word "clone" in reference to animals. Later work by Shinya Yamanaka showed how mature cells can be reprogrammed to become pluripotent, extending the possibilities to non-stem cells. Gurdon and Yamanaka were jointly awarded the Nobel Prize in 2012 for this work. [47]

Humans Edit

True polyploidy rarely occurs in humans, although polyploid cells occur in highly differentiated tissue, such as liver parenchyma, heart muscle, placenta and in bone marrow. [1] [48] Aneuploidy is more common.

Polyploidy occurs in humans in the form of triploidy, with 69 chromosomes (sometimes called 69, XXX), and tetraploidy with 92 chromosomes (sometimes called 92, XXXX). Triploidy, usually due to polyspermy, occurs in about 2–3% of all human pregnancies and

15% of miscarriages. [ citation needed ] The vast majority of triploid conceptions end as a miscarriage those that do survive to term typically die shortly after birth. In some cases, survival past birth may be extended if there is mixoploidy with both a diploid and a triploid cell population present. There has been one report of a child surviving to the age of seven months with complete triploidy syndrome. He failed to exhibit normal mental or physical neonatal development, and died from a Pneumocystis carinii infection, which indicates a weak immune system. [49]

Triploidy may be the result of either digyny (the extra haploid set is from the mother) or diandry (the extra haploid set is from the father). Diandry is mostly caused by reduplication of the paternal haploid set from a single sperm, but may also be the consequence of dispermic (two sperm) fertilization of the egg. [50] Digyny is most commonly caused by either failure of one meiotic division during oogenesis leading to a diploid oocyte or failure to extrude one polar body from the oocyte. Diandry appears to predominate among early miscarriages, while digyny predominates among triploid zygotes that survive into the fetal period. [51] However, among early miscarriages, digyny is also more common in those cases less than 8 + 1 ⁄ 2 weeks gestational age or those in which an embryo is present. There are also two distinct phenotypes in triploid placentas and fetuses that are dependent on the origin of the extra haploid set. In digyny, there is typically an asymmetric poorly grown fetus, with marked adrenal hypoplasia and a very small placenta. [ citation needed ] In diandry, a partial hydatidiform mole develops. [50] These parent-of-origin effects reflect the effects of genomic imprinting. [ citation needed ]

Complete tetraploidy is more rarely diagnosed than triploidy, but is observed in 1–2% of early miscarriages. However, some tetraploid cells are commonly found in chromosome analysis at prenatal diagnosis and these are generally considered 'harmless'. It is not clear whether these tetraploid cells simply tend to arise during in vitro cell culture or whether they are also present in placental cells in vivo. There are, at any rate, very few clinical reports of fetuses/infants diagnosed with tetraploidy mosaicism.

Mixoploidy is quite commonly observed in human preimplantation embryos and includes haploid/diploid as well as diploid/tetraploid mixed cell populations. It is unknown whether these embryos fail to implant and are therefore rarely detected in ongoing pregnancies or if there is simply a selective process favoring the diploid cells.

Fishes Edit

A polyploidy event occurred within the stem lineage of the teleost fishes. [35]

Plants Edit

Polyploidy is frequent in plants, some estimates suggesting that 30–80% of living plant species are polyploid, and many lineages show evidence of ancient polyploidy (paleopolyploidy) in their genomes. [52] [53] [54] [55] Huge explosions in angiosperm species diversity appear to have coincided with the timing of ancient genome duplications shared by many species. [56] It has been established that 15% of angiosperm and 31% of fern speciation events are accompanied by ploidy increase. [57]

Polyploid plants can arise spontaneously in nature by several mechanisms, including meiotic or mitotic failures, and fusion of unreduced (2n) gametes. [58] Both autopolyploids (e.g. potato [59] ) and allopolyploids (such as canola, wheat and cotton) can be found among both wild and domesticated plant species.

Most polyploids display novel variation or morphologies relative to their parental species, that may contribute to the processes of speciation and eco-niche exploitation. [53] [58] The mechanisms leading to novel variation in newly formed allopolyploids may include gene dosage effects (resulting from more numerous copies of genome content), the reunion of divergent gene regulatory hierarchies, chromosomal rearrangements, and epigenetic remodeling, all of which affect gene content and/or expression levels. [60] [61] [62] [63] Many of these rapid changes may contribute to reproductive isolation and speciation. However seed generated from interploidy crosses, such as between polyploids and their parent species, usually suffer from aberrant endosperm development which impairs their viability, [64] [65] thus contributing to polyploid speciation.

Some plants are triploid. As meiosis is disturbed, these plants are sterile, with all plants having the same genetic constitution: Among them, the exclusively vegetatively propagated saffron crocus (Crocus sativus). Also, the extremely rare Tasmanian shrub Lomatia tasmanica is a triploid sterile species.

There are few naturally occurring polyploid conifers. One example is the Coast Redwood Sequoia sempervirens, which is a hexaploid (6x) with 66 chromosomes (2n = 6x = 66), although the origin is unclear. [66]

Aquatic plants, especially the Monocotyledons, include a large number of polyploids. [67]

Crops Edit

The induction of polyploidy is a common technique to overcome the sterility of a hybrid species during plant breeding. For example, triticale is the hybrid of wheat (Triticum turgidum) and rye (Secale cereale). It combines sought-after characteristics of the parents, but the initial hybrids are sterile. After polyploidization, the hybrid becomes fertile and can thus be further propagated to become triticale.

In some situations, polyploid crops are preferred because they are sterile. For example, many seedless fruit varieties are seedless as a result of polyploidy. Such crops are propagated using asexual techniques, such as grafting.

Polyploidy in crop plants is most commonly induced by treating seeds with the chemical colchicine.

Examples Edit
  • Triploid crops: some apple varieties (such as Belle de Boskoop, Jonagold, Mutsu, Ribston Pippin), banana, citrus, ginger, watermelon, [68]saffron crocus, white pulp of coconut
  • Tetraploid crops: very few apple varieties, durum or macaroniwheat, cotton, potato, canola/rapeseed, leek, tobacco, peanut, kinnow, Pelargonium
  • Hexaploid crops: chrysanthemum, bread wheat, triticale, oat, kiwifruit[8]
  • Octaploid crops: strawberry, dahlia, pansies, sugar cane, oca (Oxalis tuberosa) [69]
  • Dodecaploid crops: some sugar cane hybrids [70]

Some crops are found in a variety of ploidies: tulips and lilies are commonly found as both diploid and triploid daylilies (Hemerocallis cultivars) are available as either diploid or tetraploid apples and kinnow mandarins can be diploid, triploid, or tetraploid.

Fungi Edit

Besides plants and animals, the evolutionary history of various fungal species is dotted by past and recent whole-genome duplication events (see Albertin and Marullo 2012 [71] for review). Several examples of polyploids are known:

  • autopolyploid: the aquatic fungi of genus Allomyces, [72] some Saccharomyces cerevisiae strains used in bakery, [73] etc.
  • allopolyploid: the widespread Cyathus stercoreus, [74] the allotetraploid lager yeast Saccharomyces pastorianus, [75] the allotriploid wine spoilage yeast Dekkera bruxellensis, [76] etc.
  • paleopolyploid: the human pathogen Rhizopus oryzae, [77] the genus Saccharomyces, [78] etc.

In addition, polyploidy is frequently associated with hybridization and reticulate evolution that appear to be highly prevalent in several fungal taxa. Indeed, homoploid speciation (hybrid speciation without a change in chromosome number) has been evidenced for some fungal species (such as the basidiomycota Microbotryum violaceum [79] ).

As for plants and animals, fungal hybrids and polyploids display structural and functional modifications compared to their progenitors and diploid counterparts. In particular, the structural and functional outcomes of polyploid Saccharomyces genomes strikingly reflect the evolutionary fate of plant polyploid ones. Large chromosomal rearrangements [80] leading to chimeric chromosomes [81] have been described, as well as more punctual genetic modifications such as gene loss. [82] The homoealleles of the allotetraploid yeast S. pastorianus show unequal contribution to the transcriptome. [83] Phenotypic diversification is also observed following polyploidization and/or hybridization in fungi, [84] producing the fuel for natural selection and subsequent adaptation and speciation.

Chromalveolata Edit

Other eukaryotic taxa have experienced one or more polyploidization events during their evolutionary history (see Albertin and Marullo, 2012 [71] for review). The oomycetes, which are non-true fungi members, contain several examples of paleopolyploid and polyploid species, such as within the genus Phytophthora. [85] Some species of brown algae (Fucales, Laminariales [86] and diatoms [87] ) contain apparent polyploid genomes. In the Alveolata group, the remarkable species Paramecium tetraurelia underwent three successive rounds of whole-genome duplication [88] and established itself as a major model for paleopolyploid studies.

Bacteria Edit

Each Deinococcus radiodurans bacterium contains 4-8 copies of its chromosome. [89] Exposure of D. radiodurans to X-ray irradiation or desiccation can shatter its genomes into hundred of short random fragments. Nevertheless, D. radiodurans is highly resistant to such exposures. The mechanism by which the genome is accurately restored involves RecA-mediated homologous recombination and a process referred to as extended synthesis-dependent strand annealing (SDSA). [90]

Azotobacter vinelandii can contain up to 80 chromosome copies per cell. [91] However this is only observed in fast growing cultures, whereas cultures grown in synthetic minimal media are not polyploid. [92]

Archaea Edit

The archaeon Halobacterium salinarium is polyploid [93] and, like Deinococcus radiodurans, is highly resistant to X-ray irradiation and desiccation, conditions that induce DNA double-strand breaks. [94] Although chromosomes are shattered into many fragments, complete chromosomes can be regenerated by making use of overlapping fragments. The mechanism employs single-stranded DNA binding protein and is likely homologous recombinational repair. [95]


Methods

Identification of MLO Proteins

Using Arabidopsis, tomato and rice MLO protein sequences representing all clades as a query, translated plant genomes were examined for MLO proteins using BLASTP ( Altschul et al. 1997). The following databases were used for these searches: NCBI http://blast.ncbi.nlm.nih.gov/Blast.cgi ( Altschul et al. 1997), http://solgenomics.net/ (N. benthamiana Genome v. 1.0.1), Phytozome v10.2 http://phytozome.jgi.doe.gov/pz/portal.html (A. coerulea v1.1, P. patens v3.0, S. moellendorfii v.1.0, S. italica v2.1), http://congenie.org/ (P. abies v1.0, P. taeda v1.0), http://www.amborella.org/ (Amborella genome scaffold v1.0), http://webblast.ipk-gatersleben.de/barley/ (H. vulgare v1). Database quests for non-land plant MLOs were performed in autumn 2015. Sequences were examined manually for apparent completeness and correctness against an alignment of published MLO protein sequences as reference, using BioEdit v7.1.11 ( Hall 1999).

Alignments and Phylogenetic Analysis

For further analysis, all MLO amino acid sequences were aligned using ClustalW implemented in the MEGA v6.0 software ( Tamura et al. 2013). The alignment was further optimized by manual inspection and curation. The phylogenetic trees were generated with MEGA v6.0, using the Maximum-Likelihood, Maximum parsimony, Neighbor-Joining, and UPGMA methods based on the J ones– T aylor– T hornton (JTT) matrix model, with 1,000 bootstrap replications each. The consensus sequences were generated using BioEdit v.7.1.11 ( Hall 1999). Minimal consensus sequences were generated with a threshold of 0%, local conservation levels were determined with consensus sequences using thresholds of 90% (high conservation) and 95% (quasi-invariant residues).


Results

Computing Gene Families and Flux Values

We estimated the flux through each biochemical reaction in the Arabidopsis and sorghum metabolic networks using flux-balance analysis ( Orth et al. 2010), maximizing the production of new cell mass for a fixed input of either light energy in both Arabidopsis and sorghum (in photosynthetic tissues) or carbohydrates for Arabidopsis (in nonphotosynthetic tissues, see Materials and Methods). We included the sorghum network to be sure that the differences in C3 and C4 photosynthesis were not greatly biasing our results.

Maximal flux values ranged from 0 to 3865120 (arbitrary flux-balance units) in the Arabidopsis leaf, from 0 to 6156740 in the Arabidopsis root, and from 0 to 2560860 in sorghum, when the biomass production is maximized and scaled to 1000 units. We then coupled those data to a set of cross-genome gene families identified from the 10 plant genomes (Materials and Methods). The result was a set of 735 gene families with associated metabolic fluxes. Of these 735 gene families, 463 have absolute flux values greater than zero. These families vary in size from 4 to 306 genes. The number of non–null-flux values associated with each family ranges from 1 to 13, with 90% having only one associated flux value and only three having 10 or more flux values. Those three families function as ATP synthases, phospholipid transporters, and cellulose synthases (functional Gene Ontology annotation from TAIR Swarbreck et al. 2008). The number of gene duplications per family varies from 0 to 210, with a mean of 3.21 duplications per species. Reactions with no flux can result either from failure to include certain metabolites in the biomass reaction or from a reaction not being used in certain conditions. Because of the potential for error introduced by these two possibilities, we present our results both with and without null-flux reactions.

Correlation Between Number of Duplications and Maximum Metabolic Flux

The correlation between the number of duplications in a gene family and the maximal flux is positive and significant for both C3 and C4 model networks, whether or not null-flux reactions are included and whether duplications are calculated per species or per family ( table 1).

Correlations Between Duplication and Flux by Gene Family

All Flux Values Excluding Null-Flux a
r b P c rP
Duplications per gene family
All conditions 0.245 <10 −15 0.336 <10 −15
C3 leaves 0.218 <10 −15 0.328 <10 −15
C4 leaves 0.176 <10 −8 0.218 <10 −4
Roots 0.223 <10 −15 0.359 <10 −15
Duplications per species per gene family d
All conditions 0.227 <10 −15 0.306 <10 −15
C3 leaves 0.203 <10 −15 0.272 <10 −14
C4 leaves 0.163 <10 −7 0.206 <10 −4
Roots 0.211 <10 −15 0.342 <10 −15
All Flux Values Excluding Null-Flux a
r b P c rP
Duplications per gene family
All conditions 0.245 <10 −15 0.336 <10 −15
C3 leaves 0.218 <10 −15 0.328 <10 −15
C4 leaves 0.176 <10 −8 0.218 <10 −4
Roots 0.223 <10 −15 0.359 <10 −15
Duplications per species per gene family d
All conditions 0.227 <10 −15 0.306 <10 −15
C3 leaves 0.203 <10 −15 0.272 <10 −14
C4 leaves 0.163 <10 −7 0.206 <10 −4
Roots 0.211 <10 −15 0.342 <10 −15

Flux values equaling 0 can have confounding biological and computational meanings.

Correlations and statistical significance calculated in R.

Number of duplication events per gene family divided by the number of species in that family.

Correlations Between Duplication and Flux by Gene Family

All Flux Values Excluding Null-Flux a
r b P c rP
Duplications per gene family
All conditions 0.245 <10 −15 0.336 <10 −15
C3 leaves 0.218 <10 −15 0.328 <10 −15
C4 leaves 0.176 <10 −8 0.218 <10 −4
Roots 0.223 <10 −15 0.359 <10 −15
Duplications per species per gene family d
All conditions 0.227 <10 −15 0.306 <10 −15
C3 leaves 0.203 <10 −15 0.272 <10 −14
C4 leaves 0.163 <10 −7 0.206 <10 −4
Roots 0.211 <10 −15 0.342 <10 −15
All Flux Values Excluding Null-Flux a
r b P c rP
Duplications per gene family
All conditions 0.245 <10 −15 0.336 <10 −15
C3 leaves 0.218 <10 −15 0.328 <10 −15
C4 leaves 0.176 <10 −8 0.218 <10 −4
Roots 0.223 <10 −15 0.359 <10 −15
Duplications per species per gene family d
All conditions 0.227 <10 −15 0.306 <10 −15
C3 leaves 0.203 <10 −15 0.272 <10 −14
C4 leaves 0.163 <10 −7 0.206 <10 −4
Roots 0.211 <10 −15 0.342 <10 −15

Flux values equaling 0 can have confounding biological and computational meanings.

Correlations and statistical significance calculated in R.

Number of duplication events per gene family divided by the number of species in that family.

Association of Flux and Duplication is neither Taxa nor Duplication-Mechanism Specific

As described, these species share a history of WGD ( fig. 1). We summed the number of duplications on each branch in figure 1, separating those with lineage-specific WGDs from those without. Duplications in both groups are significantly and positively correlated with maximum flux (WGD: r = 0.111, P < 0.05 SSD: r = 0.094, P < 0.05). Of course, the branches containing WGDs will also have some background level of SSD, meaning that the duplications on these branches will not be exclusively due to WGD. However, the similarity in correlations seen between the two types of branch suggests that a more careful accounting of duplicates is unlikely to yield different results. Similarly, we found significant positive associations of duplication and flux for the monocot subtree as well as the eudicot tree with A. thaliana removed (P < 0.05). The similarity of the results for these subtrees implies that our results are not specific to Arabidopsis, even though one of the primary metabolic networks used is from this organism. Among the terminal nodes with rice and soybean show significant associations of flux and duplication after a Bonferroni multiple-testing correction (P < 0.00256). Unfortunately, for the remainder of the tip taxa, it is difficult to distinguish between the lack of an association and the lack of sufficient numbers of duplicates to discern if that association might exist. Similarly, the flux values inferred from the sorghum C4 leaves show a mixed pattern of associations and lack thereof depending on the precise data set used (0.1689 ≤ P ≤ 0.9653).

Association of Flux and Duplication Extends Across Compartments and Functional Annotations

Gene families were associated with GO Slim annotations ( supplementary table 1 , Supplementary Material online) for both cellular compartment and function. We found significant Spearman's correlations between the flux and duplication rate for the metabolic gene families from the chloroplast and mitochondria ( table 2). Likewise, gene families that have a role in DNA or RNA binding or metabolism, hydrolase activity, and responses to stimuli or stress had significant correlations between the number of duplications and flux ( table 3).

Duplication Status per Gene Family Split by Cellular Compartment

Duplication versus Flux a Duplication b
Cellular Compartment nr c PZ d P
Nucleus 56 0.320 0.016 3.481 0.0005
Cytosol 74 0.133 0.258 4.910 <0.0001
Chloroplast and plastid 273 0.275 <0.0001−0.646 0.518
Mitochondria 134 0.434 <0.00011.012 0.311
Plasma membrane 97 0.135 0.187 7.371 <0.0001
Endoplasmic reticulum 44 0.304 0.045 −1.161 0.246
Golgi apparatus 12 0.401 0.196 2.280 0.023
Cell wall 52 0.343 0.013 3.210 0.001
Extracellular 51 0.089 0.532 5.115 <0.0001
Duplication versus Flux a Duplication b
Cellular Compartment nr c PZ d P
Nucleus 56 0.320 0.016 3.481 0.0005
Cytosol 74 0.133 0.258 4.910 <0.0001
Chloroplast and plastid 273 0.275 <0.0001−0.646 0.518
Mitochondria 134 0.434 <0.00011.012 0.311
Plasma membrane 97 0.135 0.187 7.371 <0.0001
Endoplasmic reticulum 44 0.304 0.045 −1.161 0.246
Golgi apparatus 12 0.401 0.196 2.280 0.023
Cell wall 52 0.343 0.013 3.210 0.001
Extracellular 51 0.089 0.532 5.115 <0.0001

Bold values are significant at a Bonferroni corrected α = 0.0055.

Duplications per gene family versus the maximum flux.

Wilcoxon rank test of difference across compartments (positive values: overduplication negative values: underduplication).

Spearman's r, calculated in SAS (v9.2.2, Cary, NC).

Wilcoxon's Z, calculated in SAS (v9.2.2, Cary, NC).

Duplication Status per Gene Family Split by Cellular Compartment

Duplication versus Flux a Duplication b
Cellular Compartment nr c PZ d P
Nucleus 56 0.320 0.016 3.481 0.0005
Cytosol 74 0.133 0.258 4.910 <0.0001
Chloroplast and plastid 273 0.275 <0.0001−0.646 0.518
Mitochondria 134 0.434 <0.00011.012 0.311
Plasma membrane 97 0.135 0.187 7.371 <0.0001
Endoplasmic reticulum 44 0.304 0.045 −1.161 0.246
Golgi apparatus 12 0.401 0.196 2.280 0.023
Cell wall 52 0.343 0.013 3.210 0.001
Extracellular 51 0.089 0.532 5.115 <0.0001
Duplication versus Flux a Duplication b
Cellular Compartment nr c PZ d P
Nucleus 56 0.320 0.016 3.481 0.0005
Cytosol 74 0.133 0.258 4.910 <0.0001
Chloroplast and plastid 273 0.275 <0.0001−0.646 0.518
Mitochondria 134 0.434 <0.00011.012 0.311
Plasma membrane 97 0.135 0.187 7.371 <0.0001
Endoplasmic reticulum 44 0.304 0.045 −1.161 0.246
Golgi apparatus 12 0.401 0.196 2.280 0.023
Cell wall 52 0.343 0.013 3.210 0.001
Extracellular 51 0.089 0.532 5.115 <0.0001

Bold values are significant at a Bonferroni corrected α = 0.0055.

Duplications per gene family versus the maximum flux.

Wilcoxon rank test of difference across compartments (positive values: overduplication negative values: underduplication).

Spearman's r, calculated in SAS (v9.2.2, Cary, NC).

Wilcoxon's Z, calculated in SAS (v9.2.2, Cary, NC).

Duplication Status per Gene Family Split by Functional Annotation

Duplication versus Flux a Duplication b
Function nr c PZ d P
Cell organization and biogenesis 29 0.209 0.274 1.010 0.312
Developmental processes 20 −0.132 0.578 1.693 0.090
DNA or RNA binding or metabolism 26 0.760 <0.0001−2.284 0.022
Electron transport 7 0.860 0.013 0.460 0.645
Hydrolase activity 114 0.310 <0.001 −1.661 0.097
Kinase activity 62 −0.031 0.817 1.167 0.243
Nucleic acid or Nucleotide binding 94 0.062 0.554 0.011 0.991
Protein binding or metabolism 121 0.139 0.128 1.463 0.143
Signal transduction 13 0.104 0.735 2.450 0.014
Stimulus or stress response 199 0.302 <0.00012.650 0.008
Transferase activity 166 0.211 0.006 −0.833 0.405
Transporters or transport 56 −0.034 0.801 1.755 0.079
Duplication versus Flux a Duplication b
Function nr c PZ d P
Cell organization and biogenesis 29 0.209 0.274 1.010 0.312
Developmental processes 20 −0.132 0.578 1.693 0.090
DNA or RNA binding or metabolism 26 0.760 <0.0001−2.284 0.022
Electron transport 7 0.860 0.013 0.460 0.645
Hydrolase activity 114 0.310 <0.001 −1.661 0.097
Kinase activity 62 −0.031 0.817 1.167 0.243
Nucleic acid or Nucleotide binding 94 0.062 0.554 0.011 0.991
Protein binding or metabolism 121 0.139 0.128 1.463 0.143
Signal transduction 13 0.104 0.735 2.450 0.014
Stimulus or stress response 199 0.302 <0.00012.650 0.008
Transferase activity 166 0.211 0.006 −0.833 0.405
Transporters or transport 56 −0.034 0.801 1.755 0.079

Bold values are significant at a Bonferroni corrected α = 0.0042.

Duplications per gene family versus the maximum flux.

Wilcoxon rank test of difference across compartments (positive values: overduplication negative values: underduplication).

Spearman's r, calculated in SAS (v9.2.2, Cary, NC).

Wilcoxon's Z, calculated in SAS (v9.2.2, Cary, NC).

Duplication Status per Gene Family Split by Functional Annotation

Duplication versus Flux a Duplication b
Function nr c PZ d P
Cell organization and biogenesis 29 0.209 0.274 1.010 0.312
Developmental processes 20 −0.132 0.578 1.693 0.090
DNA or RNA binding or metabolism 26 0.760 <0.0001−2.284 0.022
Electron transport 7 0.860 0.013 0.460 0.645
Hydrolase activity 114 0.310 <0.001 −1.661 0.097
Kinase activity 62 −0.031 0.817 1.167 0.243
Nucleic acid or Nucleotide binding 94 0.062 0.554 0.011 0.991
Protein binding or metabolism 121 0.139 0.128 1.463 0.143
Signal transduction 13 0.104 0.735 2.450 0.014
Stimulus or stress response 199 0.302 <0.00012.650 0.008
Transferase activity 166 0.211 0.006 −0.833 0.405
Transporters or transport 56 −0.034 0.801 1.755 0.079
Duplication versus Flux a Duplication b
Function nr c PZ d P
Cell organization and biogenesis 29 0.209 0.274 1.010 0.312
Developmental processes 20 −0.132 0.578 1.693 0.090
DNA or RNA binding or metabolism 26 0.760 <0.0001−2.284 0.022
Electron transport 7 0.860 0.013 0.460 0.645
Hydrolase activity 114 0.310 <0.001 −1.661 0.097
Kinase activity 62 −0.031 0.817 1.167 0.243
Nucleic acid or Nucleotide binding 94 0.062 0.554 0.011 0.991
Protein binding or metabolism 121 0.139 0.128 1.463 0.143
Signal transduction 13 0.104 0.735 2.450 0.014
Stimulus or stress response 199 0.302 <0.00012.650 0.008
Transferase activity 166 0.211 0.006 −0.833 0.405
Transporters or transport 56 −0.034 0.801 1.755 0.079

Bold values are significant at a Bonferroni corrected α = 0.0042.

Duplications per gene family versus the maximum flux.

Wilcoxon rank test of difference across compartments (positive values: overduplication negative values: underduplication).

Spearman's r, calculated in SAS (v9.2.2, Cary, NC).

Wilcoxon's Z, calculated in SAS (v9.2.2, Cary, NC).

To determine whether duplication rates differed among compartments or classes, we used Wilcoxon rank-sum test (Z-scores in tables 2 and 3). Although gene families could appear in more than one annotation group, families located in the nucleus, cytosol, plasma membrane, cell wall, and extracellular space were significantly overduplicated compared with all other gene families ( table 2). No functional categories were significantly overduplicated ( table 3).

Selection on Sequence Evolution of Ion Transporters

We chose to analyze the ion transporters because of their interesting role as potential chokepoints. In the metabolic networks used in this analysis, the gene families representing transporters have significantly higher flux than nontransporter gene families (Mann–Whitney one-tailed P < 10 −15 ). However, we found no significant correlation between the flux and duplicability among transporter gene families (P = 0.599). Therefore, we chose to look at the fine-scale differences in selection in two classes of ion transporters, phosphate and sodium. These elements have distinct roles in the growth and development of plants and hence potentially differing duplication dynamics. Phosphate transporters import an essential macronutrient, while sodium transporters primarily limit the import of potentially toxic sodium ( Rausch and Bucher 2002 Kronzucker and Britto 2011). By narrowing our focus to just these 5 gene families and limiting ourselves to the three species (A. thaliana, P. trichocarpa, and C. papaya), it is possible to manually isolate SSD and WGD events. This inference in turn allows us to assess if the strength of selection differs following WGD, SSD, and speciation.

Phosphate Transporters

Phosphate transporters in A. thaliana are divided into four gene families. These families include the high-affinity transporters (PHT1 Mudge et al. 2002 Poirier and Bucher 2002), which import ions across the plasma membrane, and the mitochondrial (PHT3 Hamel et al. 2004) and chloroplast (PHT4 Guo et al. 2008) transporters, which act in their respective organelles. Finally, low-affinity (PHT2) phosphate transporters are also localized to the chloroplast ( Versaw and Harrison 2002). We inferred gene phylogenies for the four phosphate transporter families and for one sodium transporter family (see Materials and Methods). Although the topology of phosphate transporter gene families is easily reconciled to the species tree, none of the clades contained the 4:2:1 ratio of A. thaliana to P. trichocarpa to C. papaya genes that would be expected if all transporters had been retained following the α, β, and P. trichocarpa–WGDs and no SSDs had been retained ( fig. 2a and b). The average selective constraint (Ka/Ks) for PHT gene families varies considerably from 0.076 in high-affinity transporters to 0.207 in low-affinity transporters ( table 4). The lowest Ka/Ks corresponds to the family with the largest observed number of duplications (high-affinity transporters: 19 duplications), whereas the highest Ka/Ks values correspond to the family with the fewest duplications (low-affinity transporters: 1 duplication). This observation is, however, without statistical significance. In all cases, the branches following gene duplications show significantly higher Ka/Ks than do those following speciation ( table 4 but note that the small size of the low-affinity family limits the strength of our conclusion for that family). We also investigated selective constraints associated with duplication mechanism by dividing the branches following duplications into those due to WGD and to SSD. Here, the difference in selective constraint is less clear: for the high-affinity and chloroplast phosphate transporters, the Ka/Ks values for whole-genome duplicates are not significantly different than those for SSDs. Among the mitochondrial transporters, whole-genome duplicates have significantly higher Ka/Ks than small-scale duplicates, indicating a weaker selective constraint following WGD. The counts of WGDs versus SSDs per gene family are statistically uninformative (Fisher's Exact test: P = 0.75).

Selective Constraint Estimated with Three Models of Gene Evolution for Ion Transporters of A. thaliana, C. papaya, and P. trichocarpa

PHT1–High-Affinity Phosphate Transporter PHT2–Low-Affinity Phosphate Transporter PHT3-Mitochondrial Phosphate Transporter PHT4-Chloroplast Phosphate Transporter NHX-Sodium Ion Transporter
Model Branches Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL
R_Null All 0.076 0.207 0.114 0.148 0.049
15379.5 3577.0 5898.3 24869.3 11210.1
R_Dupl Speciation 0.063 a 0.156 a 0.080 a 0.123 a 0.062 a
Duplication 0.082 a 0.415 a 0.133 a 0.249 a 0.031 a
15376.7 3570.2 5894.1 24847.0 11188.5
R_WGD Speciation 0.063 — b 0.080 a 0.123 0.061 a
WGD c 0.081 0.190 a 0.233 0.067 a
SSD d 0.085 0.112 a 0.265 0.018 a
15376.6 5891.2 24846.7 11175.3
PHT1–High-Affinity Phosphate Transporter PHT2–Low-Affinity Phosphate Transporter PHT3-Mitochondrial Phosphate Transporter PHT4-Chloroplast Phosphate Transporter NHX-Sodium Ion Transporter
Model Branches Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL
R_Null All 0.076 0.207 0.114 0.148 0.049
15379.5 3577.0 5898.3 24869.3 11210.1
R_Dupl Speciation 0.063 a 0.156 a 0.080 a 0.123 a 0.062 a
Duplication 0.082 a 0.415 a 0.133 a 0.249 a 0.031 a
15376.7 3570.2 5894.1 24847.0 11188.5
R_WGD Speciation 0.063 — b 0.080 a 0.123 0.061 a
WGD c 0.081 0.190 a 0.233 0.067 a
SSD d 0.085 0.112 a 0.265 0.018 a
15376.6 5891.2 24846.7 11175.3

Bold values indicate a significant improvement over the model immediately above at P < 0.05 nested likelihood ratio test (distributed χ 2 , P < 0.05, degrees of freedom = 1).

No small scale duplications in PHT2, so model R_Dupl is equivalent to model R_WGD.

WGD: determined by syntenic paralogy using the Plant Genome Duplication Database ( Tang, Bowers, et al. 2008).

SSD: determined either by a lack of syntenic paralogy and/or by tandem duplication status.

Selective Constraint Estimated with Three Models of Gene Evolution for Ion Transporters of A. thaliana, C. papaya, and P. trichocarpa

PHT1–High-Affinity Phosphate Transporter PHT2–Low-Affinity Phosphate Transporter PHT3-Mitochondrial Phosphate Transporter PHT4-Chloroplast Phosphate Transporter NHX-Sodium Ion Transporter
Model Branches Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL
R_Null All 0.076 0.207 0.114 0.148 0.049
15379.5 3577.0 5898.3 24869.3 11210.1
R_Dupl Speciation 0.063 a 0.156 a 0.080 a 0.123 a 0.062 a
Duplication 0.082 a 0.415 a 0.133 a 0.249 a 0.031 a
15376.7 3570.2 5894.1 24847.0 11188.5
R_WGD Speciation 0.063 — b 0.080 a 0.123 0.061 a
WGD c 0.081 0.190 a 0.233 0.067 a
SSD d 0.085 0.112 a 0.265 0.018 a
15376.6 5891.2 24846.7 11175.3
PHT1–High-Affinity Phosphate Transporter PHT2–Low-Affinity Phosphate Transporter PHT3-Mitochondrial Phosphate Transporter PHT4-Chloroplast Phosphate Transporter NHX-Sodium Ion Transporter
Model Branches Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL Ka/Ks−lnL
R_Null All 0.076 0.207 0.114 0.148 0.049
15379.5 3577.0 5898.3 24869.3 11210.1
R_Dupl Speciation 0.063 a 0.156 a 0.080 a 0.123 a 0.062 a
Duplication 0.082 a 0.415 a 0.133 a 0.249 a 0.031 a
15376.7 3570.2 5894.1 24847.0 11188.5
R_WGD Speciation 0.063 — b 0.080 a 0.123 0.061 a
WGD c 0.081 0.190 a 0.233 0.067 a
SSD d 0.085 0.112 a 0.265 0.018 a
15376.6 5891.2 24846.7 11175.3

Bold values indicate a significant improvement over the model immediately above at P < 0.05 nested likelihood ratio test (distributed χ 2 , P < 0.05, degrees of freedom = 1).

No small scale duplications in PHT2, so model R_Dupl is equivalent to model R_WGD.

WGD: determined by syntenic paralogy using the Plant Genome Duplication Database ( Tang, Bowers, et al. 2008).

SSD: determined either by a lack of syntenic paralogy and/or by tandem duplication status.

Sodium Transporters

The angiosperm sodium ion transporters (NHX) are a single gene family responsible for keeping Na + concentrations at nontoxic levels ( Rodríguez-Rosales et al. 2008). The sodium ion transporters have a lower average Ka/Ks than do any of the phosphate transporter families (0.049 versus 0.076–0.207). Curiously, among these transporters, paralogs have significantly lower Ka/Ks values than do orthologs, indicating no release in a selective constraint after duplication ( table 4). Genes duplicated by WGD seem to be under slightly less selective constraint than gene orthologs however, SSDs seem to be under considerably higher selective constraint than either.


CHAPTER 9 - Comparative Genomics in Eukaryotes

This chapter outlines the development and current status of comparative eukaryotic genomics, from the earliest studies of basic chromosome structure to the sequencing of entire genomes. In the process, a review is provided for the structure, organization, and composition of the primary eukaryotic genomes that have been sequenced thus far. Although the word “genome,” meaning the total hereditary material of an organism, was coined in 1920, the general concept of genome arose before 4th century, when Aristotle implicated blood as the heredity substance. The notions of “blood relations” and characteristics being “in one's blood” persist it is now known that the blood of mammals actually contains very little genetic material because their erythrocytes contain neither nuclei nor mitochondria. Although its roots can be traced back to the earliest chromosomal work, comparative genomics involving complete genome sequencing is a science still in its infancy. Fast-growing and full of potential, its maturation is expected to influence an increasingly broad array of biological disciplines. Already, widespread implications can be envisioned for evolutionary biology, medicine, and agriculture in some cases, these have already become reality. The large-scale comparison, and perhaps even manipulation, of genomes is a complex undertaking involving numerous empirical, analytical, and ethical issues. Both important challenges and exciting discoveries lie ahead for genome biology.


Biology chapter 24 - multiple choice

a. Conservation of synteny will hinder our ability to find agriculturally important genes in plants.

b. Arabidopsis has no commercial significance except as a model organism.

c. Sequencing of the rice genome was important because it is related to many other cereal crop plants.

d. Sequencing the genomes of beneficial microbes has already begun.

a. P. falciparum has many genes with similar function clustered together on its chromosomes.

b. P. falciparum is difficult for the immune system to target because it "cloaks" itself inside of red blood cells.

c. P. falciparum has inherited a unique subcellular organelle called an apicoplast from the chloroplast of an alga.

a. Conserved sequences important to the genetic basis of a human disease are most easily detected by comparison to closely related genomes.

b. The genomes of the mammalian relatives of humans are the best targets for discovering new treatments for human diseases.

c. Comparing the genomes of parasite and host is likely to reveal good drug targets to eliminate the parasite without harming the host.

d. A comparison of mouse and human genomes would help reveal functions for previously unidentified human genes.

a. Plants have an even greater genome size range than animals.

b. Most plants have about 30,000 to 40,000 genes.

c. The difference between genome sizes in wheat and rice can be explained by the fact that wheat is hexaploid (6n) while rice is diploid (2n).

d. In plants, gene families have relatively high copy numbers.

a. There is a strong correlation between the number of genes and genome size.

b. As the human genome was sequenced, the estimated number of genes continued to decrease.

c. Much of the extra DNA in humans is in the form of introns.

d. The pufferfish as about the same of genes as humans but many fewer introns in its coding DNA.

a. A significant amount of the non-protein-coding DNA consists of retrotransposon DNA.

b. Non-protein-coding DNA may code for RNAs that are translated into transcription factors.

c. A study of mouse RNA transcripts showed that many did not code for any known mouse protein.

d. A large part of the non-protein-coding DNA may be rich in regulatory RNA sequences.

a. In humans a single point mutation in the FOXP2 gene impairs speech and grammar.

b. FOXP2 may be involved in songbird singing and mouse vocalization.

c. The protein coded by the FOXP2 gene differs by only two amino acids in humans and chimps.

d. The FOXP2 gene is involved in the control of the neuromuscular pathway leading to complex sound formation.

a. Little genetic divergence has occurred since chimps and humans shared a common ancestor.

b. Except for about 1% of their genomes, chimp and human genes are identical.

c. Using microarrays to detect mRNA transcribed from known human genes, it was possible to show that almost identical transcription patterns existed in the brains of both chimp and human.

a.most of the coding genes are different.

b. most of the non-coding genes are different.

c. gene expression differs.

d. the genes are mostly the same but have been rearranged.

a. Premature stop codons can produce pseudogenes.

b. Missense mutations can produce pseudogenes.

c. Pseudogenes cause gene inactivation.

d. Pseudogenes have DNA sequences very similar to a functional gene.

a. Synteny refers to the conservation of gene order along chromosomes.

b. Synteny refers to the constancy of chromosome numbers in related clades.

c. Synteny results from polyploidization events.

d. Synteny refers to the rearrangement of gene order due to inversions.

a. Most of the foreign DNA in the human genome is ancient.

b. Most of the foreign DNA in the human genome exists as transposons.

c. Like the Drosophila genome, the human genome is constantly eliminating its foreign DNA.

a. Horizontal gene transfer is also called lateral gene transfer.

b. Horizontal gene transfer involves hitchhiking genes from other species.

c. Horizontal gene transfer was common early in life, but is absent today.

d. Gene swapping is evident in the human genome.

b. conservation of synteny

d. humans have one more chromosome than the other great apes

a. Duplicate genes can lose their ancestral function through subsequent mutation.

b. Duplicate genes can gain a derived function through subsequent mutation.

c. Duplicate genes can share the ancestral function of the original gene.

d. Gene duplication is most likely to occur in growth and development genes, immune system genes, and cell-surface receptor genes.

a. Jumping of transposons is most common in the first few generations following a polyploidization event.

b. Genome size in plants is largely determined by polyploidization events.

c. Genome downsizing following allopolyploidy usually affects the participating hybrids unequally.

d. Genome downsizing following allopolyploidy results mainly from duplicate gene loss.

a. hybridization, chromosome doubling, duplicate gene loss

b. hybridization, duplicate gene loss, chromosome doubling

c. chromosome doubling, hybridization duplicate gene loss

a. Plants, animals, and fungi share most of the same genes for intermediary metabolism, genome replication, and protein synthesis.

b. "Plant" genes include those coding for photosynthetic pathways and morphology.

c. Plants generally have larger genomes than animals and fungi.

d. Rice has fewer genes than humans.

a. the mutation rate differs in different species.

b. exposure to radiation and mutagens differs in different species.

c. the generation time differs in different species.

d. selection pressures vary in different species.

a. A comparison of genomes confirms that humans and chimpanzees are sibling species.

b. Very few mutations seen in the two genomes occur in coding DNA.

c. Some insertion-deletions (indels) lead to loss of function changes in the two genomes.

d. More similarity exists between the genomes of human and chimpanzee than between human and mouse.

a. Human and mouse have about the same number of genes.

b. The human genome shares 99% of its genes with the mouse.

c. A comparison of genomes confirms that mouse and humans shared a common ancestor more recently than humans and pufferfish.

d. Conservation of genes has been great in the two genomes.

a. Genes regulating the basic cellular metabolism are conserved in both human and pufferfish.

b. About 25% of human genes have no counterpart in the pufferfish genome.

c. Being ancestral to humans, the pufferfish genome has more repetitive DNA than the human genome.

d. Overall, the pufferfish genome has less DNA than the human genome.

a. Orthologs are likely to have the same function.

b. Both orthologs and paralogs result from gene duplication.

c. The sequence of an ortholog is more likely to be conserved.

d. Paralogs are more likely to be pseudogenes than orthologs.

a. has no foreign DNA, it is excised by DNAases when it occurs.

b. has a very small amount of foreign DNA, mostly in the end caps (telomeres) of chromosomes.

c. has a lot of foreign DNA, mostly in the end caps (telomeres) of chromosomes.

a. illegal and irresponsible.

b. impossible, but it happened frequently in the distant past.

c. infrequent but possible it happened more often in the distant past.

d. much more frequent than in the distant past.

a. DNA sequences similar to functional genes, but do not produce functionalproducts as far as we can tell.

b. DNA sequences produced in the laboratory and artificially inserted into a genome to investigate their function.

c. duplicate genes that are on the wrong chromosome but still produce the same gene product as the original gene.

d. genes that have been inserted from a different species, such as by a retrovirus, and may or may not produce a functional product in the new species.

a. have mutated dramatically away from the same genes in other primates.

b. have been inactivated, reducing our olfactory capabilities compared to other primates.

c. have been activated, enhancing our olfactory sense compared to other primates.

d. have duplicated more frequently, resulting in increased paralogs compared to other primates.


Table S1 and Figures S1–S3 can be found online in Appendix S1. Supporting data are provided as Appendices S2, S3 and S4. The raw data sets are available on Dryad (https://doi.org/10.5061/dryad.p1j7n43).

Q.H. contributed to literature search, led data extraction, and performed data analysis T.G.L. contributed to study design, literature search and data extraction, performed data analysis and contributed to manuscript writing M.R. contributed to study design, literature search and data analysis D.B. designed and supervised the study, contributed to literature search and data extraction, developed analytical tools, performed data analysis and wrote the paper with feedback from all co-authors.

Filename Description
mec14699-sup-0001-AppendixS1.pdfPDF document, 570.6 KB
mec14699-sup-0002-AppendixS2.txtplain text document, 51.4 KB
mec14699-sup-0003-AppendixS3.txtplain text document, 38.2 KB
mec14699-sup-0004-AppendixS4.txtplain text document, 38.4 KB

Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.


The authors declare no conflicts of interest.

Filename Description
tpj13979-sup-0001-FigS1.pngPNG image, 109 KB Figure S1. Dissimilarity tree of features in tomato.
tpj13979-sup-0002-FigS2.pdfPDF document, 6.1 MB Figure S2. Histograms of features.
tpj13979-sup-0003-FigS3.pngPNG image, 133.4 KB Figure S3. Arabidopsis recombination rate for CO regions and random regions.
tpj13979-sup-0004-FigS4.pdfPDF document, 1.8 MB Figure S4. Features used in the random forest model in four species.
tpj13979-sup-0005-TableS1.xlsxMS Excel, 5.6 KB Table S1. Genome information of the studied species.
tpj13979-sup-0006-TableS2.xlsxMS Excel, 5.5 KB Table S2. Student's t-test results of tomato features.
tpj13979-sup-0007-TableS3.xlsxMS Excel, 6.3 KB Table S3. Feature importance values based on random forest in tomato.
tpj13979-sup-0008-TableS4.xlsxMS Excel, 5.3 KB Table S4. Cluster membership of the features in tomato.
tpj13979-sup-0009-TableS5.xlsxMS Excel, 15.8 KB Table S5. Feature means in random and positive regions.
tpj13979-sup-0010-TableS6.xlsxMS Excel, 27.3 KB Table S6. P values of Student's t-test on the features in maize, rice, Arabidopsis and tomato.
tpj13979-sup-0011-Legends.docxWord document, 12.6 KB

Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.