Download The causes and molecular consequences of polyploidy

Ann. N.Y. Acad. Sci. ISSN 0077-8923
A N N A L S O F T H E N E W Y O R K A C A D E M Y O F SC I E N C E S
Issue: The Year in Evolutionary Biology
The causes and molecular consequences of polyploidy
in flowering plants
Gaurav D. Moghe1,2 and Shin-Han Shiu1,2,3
1
Programs in Genetics, 2 Quantitative Biology, and 3 Department of Plant Biology, Michigan State University, East Lansing,
Michigan
Address for correspondence: Shin-Han Shiu, Department of Plant Biology, Michigan State University, 2265 Molecular Plant
Sciences Building, East Lansing, MI 48824. [email protected]
Polyploidy is an important force shaping plant genomes. All flowering plants are descendants of an ancestral polyploid
species, and up to 70% of extant vascular plant species are believed to be recent polyploids. Over the past century,
a significant body of knowledge has accumulated regarding the prevalence and ecology of polyploid plants. In this
review, we summarize our current understanding of the causes and molecular consequences of polyploidization in
angiosperms. We also provide a discussion on the relationships between polyploidy and adaptation and suggest areas
where further research may provide a better understanding of polyploidy.
Keywords: whole-genome duplication; plants; adaptation; expression divergence; fractionation; molecular consequences of polyploidy
Introduction
Polyploidization results in multiplication of the
genome and an increase in gene content that frequently leads to morphological and physiological
differences between polyploids and their diploid
progenitors.1 Polyploidy is widespread among flowering plants2,3 and has been postulated as an answer
to Darwin’s “abominable mystery” regarding the
causes behind the rapid acceleration in the diversification of angiosperms.4,5 It is also a major route for
origination of new genes via gene duplication and
subsequent diversification.6,7 Although we have a
fairly good understanding about the extent of polyploidy in eukaryotes8–10 and the modes of diversification of duplicate genes derived from polyploidy,11
there is still a considerable debate about whether
polyploidy indeed confers an evolutionary advantage to the organism and if it does, whether it contributes to speciation. In addition, although the primary pathways of polyploid generation have been
known for some time,12 only recently have we begun to identify the molecular consequences of polyploidization.
In this review, we first focus on the genetic and
environmental factors that influence the rates of
polyploidization. Second, we discuss the impact of
polyploidization at the molecular level. Third, we
summarize recent studies on the impact of polyploidy on morphology, physiology, and stress biology. Finally, we discuss current evidence on how
polyploidy contributes to adaptation and speciation. Our goal in this review is to present a brief
overview of our current state of understanding regarding a few different aspects of polyploidy. For additional information, we refer the reader to several
excellent resources2,3,13–17 that have covered these
topics in greater detail.
Causes of polyploidization in flowering
plants
Cytological pathways leading to
polyploidization
Diploids mostly propagate by producing haploid gametes, which combine to produce diploid progeny
(Fig. 1A). In rare cases, polyploids can arise through
the somatic doubling of chromosomes in the zygote
(Fig. 1B) or through the production of unreduced
gametes (Fig. 1C–E). The primary mechanism for
polyploid generation is thought to be the latter.12,18
Theoretical models considering unreduced gamete
doi: 10.1111/nyas.12466
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A
B
C
Parents
2X
2X
2X
2X
2X
2X
Gametes
X
X
X
X
2X
2X
F1
2X
2X
Unreduced
gamete
formation
4X
Zygote
somatic
doubling
4X
E
D
Parents
2X
2X
2X
2X
Gametes
X
X
2X
X
F1
F1
gametes
3X
2X
AP
X
2X
AP
Gamete 1
3X
X 2X 3X
Gamete 2
Gamete 2
X 2X 3X
2X 3X 4X
2X
Gamete 1
X 2X
F2
X
X 2X 3X 4X
2X 3X 4X 5X
3X 4X 5X 6X
Figure 1. Pathways of tetraploid formation from diploid
plants. The symbol X represents the base chromosome number of the species, with 1X corresponding to haploid gametes.
(A) The normal pathway wherein a diploid is produced as F1
progeny of two diploid parents. (B) Somatic doubling leading
to tetraploid (4X) generation from diploid. (C) Fusion of unreduced gametes can lead to tetraploid generation in one step. (D)
A diploid produced in F1 may generate a certain proportion
of aneuploid gametes (AP, most of which are not viable) and
unreduced gametes that can lead to tetraploid generation in F2.
The frequency of unreduced gamete formation can be high if
the parents belong to different species and F1 is a hybrid. (E)
The triploid bridge scenario where an intermediate triploid produces unreduced gametes leading to generation of tetraploids
and individuals of higher ploidy. These pathways of tetraploid
formation have been adapted from information presented in
Ref. 12. For examples of polypoids formed via each pathway,
as well as pathways of formation of polyploids of higher ploidy
levels, please refer to the original publication.
formation and fertilities of plants with different
ploidy levels have been used to predict equilibrium
ploidy frequencies.19–21 Notably, to fit the existing data on ploidy frequency observed in multiple
autopolyploid species, the unreduced gamete frequency was estimated to be 0.89%.21 This high rate
of unreduced gamete production is consistent with
its involvement in angiosperm polyploidy.
2
Unreduced gametes can be formed in three different ways: (1) premeiotic genome doubling due
to endoreplication mechanisms, including endocycling (alternating periods of S phase, where DNA is
replicated, and gap phase, without cell division), endomitosis (mitosis without the final cell division),22
or nuclear fusion; (2) impairments in meiosis, which
can affect either the first or the second meiotic divisions; and (3) postmeiotic genome doubling.12 The
unreduced gametes from diploids (2X, with X being
the base chromosome number of the species) can
lead to a tetraploid (4X) in one step by hybridization between unreduced male and female gametes
(type I pathway (Fig. 1C–D)) or through the creation of an intermediate triploid (type II pathway
(Fig. 1E)).19 Given that unreduced gametes can be
produced at an appreciable frequency,21 it is conceivable that unreduced gametes generated in two
individuals, or from the same individual (if selfing
is feasible), may hybridize and generate polyploids
through the type I pathway.
In the type II pathway, an unreduced gamete hybridizes with a normal gamete to produce a triploid
plant (3X). The triploid produces mostly aneuploid
gametes, which are generally not viable, and a small
percentage of viable X, 2X, or 3X gametes. These
gametes can then hybridize with other X, 2X, or 3X
gametes to generate plants of higher ploidy levels
(Fig. 1E). Hence, triploid plants are regarded as a
bridge toward polyploidy (triploid bridge), rather
than a dead end (triploid block).12 Currently, it remains unclear which pathway is more prevalent.13
Although the production of 3X gametes required in
the type II pathway would be rare in nature, unreduced gametes in artificially generated hybrids of
multiple Brassica species are produced at a much
higher frequency than in their parents. These gametes have a size distribution corresponding to >2X
genome complement, and they are more viable than
reduced gametes in the Brassica hybrids. These results support the hypothesis that the triploid bridge
scenario may be more prevalent for polyploids generated from interspecific hybridridization.12
Genetic components contributing to
polyploidization
Several Arabidopsis thaliana genes that can influence the frequency of unreduced gamete formation have been identified.23,24 For example, 60%
of the seeds produced from a mutated version of the
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SWI1/DYAD protein are triploid.25 SWI1/DYAD is
required for a proper meiosis I in both male and
female germ cells. In the SWI1/DYAD mutant, cells
skip the reduction division in meiosis I and directly
advance to the equatorial division in meiosis II, producing predominantly unreduced gametes.26,27 Mutations in several other genes lead to the production
of unreduced gametes by affecting different meiotic
and mitotic steps. For example, mutations in the
gene for the GLUCAN SYNTHASE LIKE 8 protein,
which lays down the glucan chains at cell plates during cell division, lead to a flower containing both
diploid and polyploid somatic cells. The polyploid
cells then go on to produce unreduced gametes.28
As discussed earlier, one important step in the
pathway to polyploidy is the formation of triploid
intermediates. The major challenge upon forming
this intermediate is the triploid block, originally described as the difficulty in generating viable triploids
through diploid–tetraploid crosses,29 which can
lead to reproductive isolation of the newly formed
polyploid owing to minority cytotype exclusion.30
A recent study demonstrated that one genetic component of the triploid block in A. thaliana is the
paternally expressed gene ADMETOS.31 This study
involved a mutant screen in a JASON mutant background, which produces unreduced gametes at a
high frequency, but 30% of the triploid seeds produced are aborted. The ADMETOS-1 JASON double mutant, on the other hand, has only 2% aborted
triploid seeds. The ADMETOS-1 mutant is a gainof-function mutant with elevated expression of the
ADMETOS gene.31 These results suggest that there is
genetic control over triploid formation. While such
genetic control may exist to create a postzygotic reproductive barrier for gene flow between species,
naturally occurring variation in such control mechanisms may provide an opportunity for polyploids
to be generated.
On the basis of the finding that unreduced gamete
formation is a trait with high heritability (e.g., 0.40–
0.60 in alfalfa and clover32,33 ), at least in domesticated crops experiencing artificial selection, polyploid formation through unreduced gamete and
triploid bridge formation is expected to have a significant genetic component. Although these genetic
studies are highly informative, it remains unclear
whether these newly identified genes are involved in
increasing or decreasing the rate of polyploid formation in nature. If these genes are the targets of
Plant polyploidy
selection for polyploidization frequency, they may
display substantial variation between plant species
and/or populations that have variable relative abundances of individuals with different ploidy levels.
Relationship between environment and
polyploidy
Nearly 80 years ago, it was demonstrated that a correlation exists between polyploidy and latitudinal
cline,34 suggesting potential habitat differentiation
between plants with different ploidy levels due to the
differences in latitudinal environment. In addition,
it was shown in 1920 that hot water–treated Pisum
root tips have increased frequency of tetraploidy in
somatic cells.35 In 1932, Randolf demonstrated that
high temperature (47–48 °C) results in an increased
frequency of tetraploid embryos in maize.36 Subsequently, a number of studies have established that
temperature stress, herbivory, pathogen attack, nutritional stress, and water stress, lead to an elevated
rate of unreduced gamete production.12
In addition to observations linking environmental stress, polyploidy, and unreduced gamete production, a large number of studies have focused on
how environment influences chromosome behavior and unreduced gamete production.18 Although
unreduced gametes can be produced due to premeiotic, meiotic, or postmeiotic aberrations, recent studies have shown that environmentally induced production of unreduced pollen is mainly
due to meiotic irregularities, particularly during
telophase II. In Rosa species, the proportion of unreduced pollen produced due to elevated temperature
(36 °C) differs greatly at different microspore
stages.37 The elevated temperature led to formation
of normal rose pollen tetrads as well as abnormal
dyads, triads, and polyads as a result of misorientation of meiotic spindles.
Although similarly affected during telophase II,
the formation of A. thaliana unreduced pollen due
to cold shock (4–5 °C) is not attributable to defects in spindle fibers attached to the chromosomes
but mainly to abnormalities in equatorial cell plate
formation as a result of misplaced microtubules.28
Because the aberration in cell plate formation occurs during telophase II, the cell plate separating
homologous chromosomes is in place but the one
separating sister chromatids is defective. Therefore,
instead of forming pollen tetrads (1X), 2X dyads are
formed, each containing two sets of exactly identical
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chromosomes. It is not clear if the differences in the
mechanistic details (spindles vs. cell plate formation) are due to the differences in the type of stress
applied or to differences between species.
In this section, we reviewed studies focused on
finding the genetic and environmental causes of
polyploidization, most of which act by affecting
mitosis or meiosis, thus producing unreduced gametes. A polyploid, once created, has to establish itself, and the process of neopolyploid establishment continues in the backdrop of molecular
and physiological changes occurring as a result of
genome duplication (and merging two different
genomes in allopolyploids) that a neopolyploid has
to undergo.1 In the remainder of the review, we focus on the impact of polyploidy on genome content,
on gene expression, on morphology, and, finally, on
adaptation.
Impact of polyploidy on genome content
Changes in genome organization
Polyploids have a tendency to return to a diploidized
state over time, experiencing changes in chromosome organization, gene order, expression, epigenetic modification, and biological network topology, a phenomenon known as diploidization.38
Diploidization may begin with large-scale changes
in the genome of neopolyploid plants, such as abnormal chromosome segregation, rearrangement,
and breakage,39,40 and may occur in a haphazard
manner after polyploidization41 (Fig. 2A–B). For example, in synthetic allotetraploids between doublehaploid Brassica oleracea (C genome) and Brassica
rapa (A genome), chromosomal segregation aberrations lead to extensive aneuploidy as early as the first
generation, when the aneuploidy rate is 24%.42 This
rate rises to 95% in the 11th generation. Despite the
high rate of aneuploidy, the number of homeologs
for a particular chromosome is frequently maintained at four copies (i.e., the loss of chromosome 1
from the A genome is usually associated with gain
of the same chromosome from the C genome, and
vice versa). This compensating aneuploidy suggests
a dosage balance requirement, at least in the early
generations. Compensating aneuploidy also occurs
in the naturally occurring allotetraploid Tragopogon
miscellus, in which 85% of aneuploid plants were
found to have the expected chromosome number.43
Chromosomal losses in early generations have also
4
A
P1
Chromosome loss
P2
Chr A
Translocation
t
Chr B
Chromosomal fragment loss
B
P1
P2
t
Homeologous
genes
Gene losses
C
Transposable element
P1
P2
Homeologous
genes
t
Proliferation
Transcriptional
activation or
repression
Insertion into gene
Figure 2. Genomic consequences of polyploidy. (A) Some possible scenarios with respect to genomic rearrangements, such
as chromosome loss, chromosomal translocation, and chromosome fragment loss, have been depicted in a simplified manner
using only two chromosomes. P1, parent 1; P2, parent 2. (B)
The process of gene loss in a parent-of-origin manner, termed
fractionation. In the depicted scenario, the chromosomal copy
from P2 loses most of the genes. (C) Proliferation of transposable elements over time. Such proliferation may lead to changes
in gene order, gene function, and gene expression.
been reported for synthetic allohexaploids (Triticum
aestivum,44 Brassica carinata × B. rapa,44 A. thaliana
× Arabidopsis suecica45 ) as well as autopolyploid
potato,46 alfalfa, and corn.47 Recently, the cause of
such chromosomal losses, which occur as the result
of meiotic instabilities, was tracked down to a single
locus called BOY NAMED SUE (BYS) in synthetic allopolyploids of A. thaliana × Arabidopsis arenosa.48
The authors speculate that the BYS locus may play a
role in A. suecica, which is a naturally occurring allopolyploid of A. thaliana × A. arenosa, in ensuring
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that homeologous chromosomes do not pair with
each other, a process that may lead to chromosomal
dosage irregularities in the progeny.48
In addition to changes in chromosome numbers,
newly generated polyploids display an elevated rate
of genome rearrangements leading to loss of chromosomal fragments (Fig. 2A). By tracking a limited number of markers, synthetic autotetraploids
of Paspalum notatum49 and Elymus elongatus50 were
shown to lose 10% of genome sequence in the first
generation. In Phlox drumondii, up to 25% reduction in parental DNA content was observed as early
as the third generation.50 On the contrary, studies in
synthetic A. thaliana autopolyploids reveal little to
no loss.51 These observations suggest that genome
rearrangement can be prominent in allo- and some
autopolyploids.
Duplicate gene loss and retention
Polyploidization initially results in multiplication
of gene content; however, the predominant fate
of gene duplicates is loss.52 Studies of newly sequenced genomes shed light on the extent of
gene loss in species that underwent polyploidization events several million years ago (Ma). In the
A. thaliana genome, only 17% of duplicates were
retained after a paleopolyplodization (␤) event that
took place 50 Ma.53 In the paleopolyploid Glycine
max, two rounds of whole-genome duplications
took place 59 and 13 Ma.54 In the homologous genes from the more recent duplication event,
56.6% of duplicates are no longer detectable, compared to 74.1% genes lost after the older Glycine
polyploidization. Thus, the rates of gene loss are
4.4% and 1.3% per million years (Myr) for the
younger and the older duplication event, respectively, indicating that gene loss rate was high initially but slowed down over time.54 In B. rapa
and Raphanus raphanistrum, which experienced
genome triplication 25 Ma,55–57 assuming the ancestral gene number before triplication was similar
to that in A. thaliana (30,000), the number of extant B. rapa genes (41,000) and R. raphanistrum
genes (38,000) indicates that as many as 55% of
the genes derived from genome triplication were
lost.58,59
The process of loss of polyploidy-derived genes
is referred to as fractionation, a collection of mutational mechanisms leading to the removal of duplicates derived from polyploidization60,61 (Fig. 2B).
Plant polyploidy
Studies of gene collinearity between duplicate regions in A. thaliana,62 Z. mays,63 and B. rapa64 suggest a bias in the genes lost from certain parental
genomes. In B. rapa, one of the three subgenomes
experienced significantly fewer gene losses than
the others.58,65 This phenomenon is also reflected
at the expression level, where genes located on
one subgenome tend to have higher expression
than others, indicating genome dominance.66 The
duplicated gene copy producing the most RNA
molecules appears to be the one retained.63 It has
recently been suggested that transposon silencing by small RNAs may contribute to the phenomenon of genome dominance, with the parental
genome having the lowest proportion of transposons being the more dominant.66 Fractionation
of genes also leads to preferential gene retention,
which has been reviewed recently.61 Retained duplicates derived from polyploidization have a number of distinguishing characteristics compared to
genes that remain single copy, including biased
gene function,67,68 higher gene complexity (number of exons and protein domains),69,70 higher levels of gene expression,71 significant parental genome
dominance,63,72 and higher network connectivity.62
Duplicate genes playing a role in stress response,
development, signaling, and transcriptional regulation tend to be retained, a feature consistent
across multiple polyploidization events and time
scales.59,70
Why are duplicates with these types of characteristics retained? Retained duplicates may
experience a brief period of complete functional
redundancy but eventually obtain new functions,6
and/or partition ancestral functions leading to
subfunctionalization.73 In addition to these mechanisms, the retention of duplicate genes may be attributable to balanced gene drive/gene balance,74,75
functional buffering,69 dosage selection,76 and
escape from adaptive conflict77 (reviewed by Innan
and Kondrashov11 and Edger and Pires78 ). Among
the mechanisms explaining duplicate retention,
some imply adaptive evolution (e.g., neofunctionalization, dosage selection) while the others
require purifying selection to maintain the ancestral functions (e.g., subfunctionalization, dosage
balance). Examples of adaptive duplication are
accumulating, but it remains unclear what fraction
of gene duplicates are fixed as the result of adaptive
evolution.79
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Moghe & Shiu
Mutation and transposable element activities
Because more than one gene copy is present, increased ploidy can mask the effect of deleterious
mutations.80 Meanwhile, the newly formed polyploid species has a very small effective population size, assuming postzygotic isolation from its
parental diploids. In this situation, genetic drift
is expected to play a more dominant role in
polyploid evolution. The selective pressure against
any mutation in polyploid genomes would be
more relaxed, leading to increased frequencies of
otherwise deleterious alleles. Although it is not
clear whether the spontaneous mutation rate is
higher in polyploids compared to diploids, there is
generally a higher mutation density in mutagenized
polyploids compared to diploids.81 Because this pattern is similar between natural and synthetic polyploids, and because the mutations tend to exist in
heterozygous states, the elevated mutation density is
likely a consequence of masking recessive deleterious mutations.81 In a comparison between wheat
T. aestivum, which experienced recent wholegenome duplication, and three other nonduplicated
grass species, there are more nonsynonymous substitutions, gene structural rearrangements, and alternative splice forms of genes in wheat,82 suggesting
relaxed selection.
The elevated mutation rate in polyploids may
also be attributable to elevated transposable element activities.83,84 The proliferation of transposons
in polyploids is expected owing to reduced population size, masked deleterious transposon insertion, and/or conflict in transposition repressors due
to genome merger84 (Fig. 2C). Despite this expectation, current studies present conflicting results
regarding whether proliferation of transposons is
correlated with polyploidy.83,84 For example, the
numbers of transposons in the Au short interspersed
nuclear element (SINE) family in natural polyploid
wheat species are significantly higher than those
in diploids,85 although it is difficult to ascertain
whether the observed difference is due to polyploidy, hybridization, and/or lineage divergence.
In addition, one of three synthetic allopolyploids
has a higher number of Au SINEs by the fourth
generation.85 In this case, it remains unclear whether
genome doubling or hybridization contributes to
the higher number of Au SINEs observed.
Studies on the activities of transposons in autopolyploids are also contradictory. In A. thaliana
6
synthetic autotetraploids, activation of Sunfish, a
DNA transposon, was observed.39 On the other
hand, a study assaying naturally occurring autotetraploid A. arenosa accessions found evidence for
purifying selection against expansion of the Ac-III
transposon family.86 In addition to the possibility
of transposon proliferation, transposons may be involved in recombination events leading to sequence
losses in polyploid genomes.87–89 For example, illegitimate recombination mediated by TE elements
was shown to underlie the variation observed between diploid and polyploid wheat species in the
HARDNESS locus.90 Such transposon-mediated recombination can also contribute to differential expansion and contraction of subgenomes, as shown
in maize.91
Impact of polyploidy on gene expression
and biological networks
Expression divergence between homeologs
In addition to experiencing widespread changes at
the DNA level, polyploids have considerable differences in gene expression compared to diploids, and
this has been reviewed extensively.92–96 Divergence
in the expression states of duplicated genes may lead
to the following outcomes: they may gain new expression states (neo-functionalization6 ), partition
their ancestral functions (subfunctionalization73 ),
or lose their expression state completely, leading to eventual pseudogenization.52 In A. thaliana,
57% and 75% of the duplicates derived from the
more recent ␣ and the older ␤ polyploidization
events, respectively, were found to have diverged
in expression.67 Whole-genome duplicates tend to
diverge in expression at a slower rate than tandem
duplicates, presumably because entire intergenic regions are duplicated during polyploidization, but
only a fraction might end up being duplicated during tandem duplication owing to the random nature
of DNA breakage and recombination.97
In allopolyploids, the combination of gene sets
in two species is expected to create a transcriptome shock, defined as abrupt and rapid changes
in patterns of parental gene expression in the
polyploids.98 Through a number of intriguing studies in the past decade or so, several basic features
have emerged with regard to homeolog expression in allopolyploids. The transcriptome shock
contributes to significant differences in expression levels between homeologs in allopolyploids
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Contribution of hybridization and genome
doubling to transcriptome shock
In addition to homeolog expression bias and expression level dominance, another focus is on
the relative contribution of genome doubling and
Homeologous
genes
A
Expression level
P1
2
1
2
3
4
2
0
P2
1
2
3
4
1
1
0
Genes
Genes
Polyploidization
Divergence
Expression
level dominance
Homeolog
expression bias
2
Expression level
(albeit to varying extents94 ) and the corresponding genes in diploid parents have been documented
in Arabidopsis,99 Brassica napus,100 and cotton.101
However, a recent study also showed that, in wheat
synthetic polyploids, fewer than 1% of genes show
nonadditive expression.102 In addition, concerns on
how nonadditivity is defined in polyploids have
been raised.103 Nonetheless, analysis of nonadditive
patterns of expression in allopolyploid homeologs
has led to the discovery of two related phenomena, homeolog expression bias and expression level
dominance104 (Fig. 3A).
Homeolog expression bias takes place when
one homeolog has significantly reduced level of
expression or is silenced altogether in the
allopolyploid.105 For example, in the first RNA sequencing study examining expression bias in cotton
polyploids, 59–62% and 48% of genes are differentially expressed when comparing diploids against
the natural and synthetic polyploids, respectively.106
It was found that genes from a particular subgenome
were expressed in allopolyploids, and the nature
of the subgenome differed in natural allopolyploids versus synthetic allopolyploids.106 Interestingly, among over 25,000 cotton genes containing single nucleotide polymorphisms distinguishing
homeolog origins, only 0.71–0.75% of genes have no
detectable expression in synthetic polyploids. Given
that the cotton allopolyploidization event took place
1–2 Ma,107 there may have been insufficient time for
gene loss and/or the expression of pseudo-genized
copies are still detectable. Related to homeolog
expression bias, expression level dominance (originally called genomic dominance) describes a situation where the sum of expression levels of a homeologous gene pair tends to be more similar to that in
one parent, regardless of the expression level of the
gene in the parent in question.104,108 These phenomena have been well documented in multiple natural
and synthetic allopolyploids. The cause of expression biases may be partially attributed to cis and
trans regulatory differences between the hybridized
genomes109 and epigenetic regulation, which is discussed in a later section.
Plant polyploidy
1
0
2
1
2
3
4
1
2
3
4
1
0
Genes
DNA
methylation
B
Gene
Polyploidization
Demethylation
RNA
polymerase
Gene
Transcribed
mRNA
Repressor
C
Gene
RNA
polymerase
Polyploidization
small RNA transcription
Epigenetic remodeling
Gene
Transcribed
mRNA
small RNAs
Figure 3. Effects of polyploidy on gene expression and epigenetic regulation. (A) A hypothetical scenario depicting expression divergence upon polyploidization. In “Homeolog expression bias,” the homeologous genes are expressed in a parentof-origin manner. In “Expression level dominance,” the sum
of the expression level of both the genes is similar to that in
one parent. P1, parent 1; P2, parent 2. (B) A gene that is silenced by DNA methylation in the parent is demethylated upon
polyploidization or hybridization, leading to its transcription
by RNA polymerase. (C) A gene with a repressor bound to its
promoter region is not expressed in the parent, but upon polyploidization is expressed as the repressor is removed owing to
regulation by small RNAs transcribed elsewhere in the genome.
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Moghe & Shiu
hybridization to nonadditive expression in allopolyploids. Through comparisons of hybrid diploids,
synthetic allopolyploids, and neopolyploids (particularly Senecio and Tragopogon species, which formed
in the past 100 years110,111 ), the effect of hybridization and genome doubling on gene expression
changes has been quantified, and these studies suggest that hybridization likely plays a more dominant
role. For example, only 88 genes are differentially
expressed between A. thaliana diploid sand synthetic isogenic autotetraploids, compared to >1700
genes with significantly different expression levels between synthetic allotetraploid A. thaliana ×
A. arenosa and the average of two parents.112 In a
study of diploid, autotetraploid, and autohexaploid
Helianthus decapetalus, ploidy level does not contribute significantly to expression differences.113
Similarly in Senecio, twice as many genes in S. ×
baxteri, a triploid hybrid between S. vulgaris (2n =
4X) and S. squalidus (2n = 2X), are differentially
expressed compared to the parental species than in
a synthetic allohexaploid derived from the triploid
S. × baxteri.98 Interestingly, genome doubling ameliorates the effect of hybridization in Senecio,98
a finding that is also reported in neopolyploid
Spartina.114 Thus, the nonadditive gene expression changes in these allopolyploids is likely attributable to interspecific hybridization and not
simply genome doubling.
Although hybridization seems to play a dominant role, studies in cotton, Spartina, and domesticated rice subspecies suggest the contribution of
genome doubling may be important. In the natural polyploid cotton, among assayed genes with
expression bias toward one parental diploid, only
25% are biased in the same direction as the diploid
hybrid,101 suggesting the expression bias in the remaining 75% is the result of genome doubling.
However, allopolyploid cotton was established 1–
2 Ma. Thus, one cannot rule out the possibility that
the expression bias is due to regulatory variation
that accumulated in the past 1–2 Myr. In Spartina,
the allopolyploid tends to have a higher number of
transgressively overexpressed genes compared to the
species hybrid,114 again suggesting a prominent role
of genome doubling. Nonetheless, given that the
parental Spartina species are hexaploids, it remains
unclear if the findings are applicable to comparisons between lower ploidy levels. Finally, genome
doubling seems to have a more prominent role in
8
transcriptome shock than hybridization in a comparison between a hybrid of Oryza sativa spp. indica and japonica and synthetic tetraploid rice.109
But given that there has been repeated hybridization between japonica and indica,115 it remains unclear how hybridization may have influenced the
findings.
Involvement of epigenetic modifications and
small RNAs in transcriptome shock
Transcriptome shock results in nonadditive gene expression and silencing of homeologs. In allopolyploids, trans factors and cis regulatory components from two genetic backgrounds can interact
owing to the hybridization between two genomes,
contributing to changes in gene expression between
polyploids and their diploid progenitors.116,117 In
addition, epigenetic factors, including DNA methylation, histone modifications, and small RNA, have
been implicated in modulating gene expression
in allpolyploids.66,116,118–122 Extensive DNA methylation changes have been reported between allopolyploids and their parents in B. napus,123
wheat,124 Spartina anglica,88,125 and Arabidopsis126
(Fig. 3B). Inconsistent with the above observations, the synthetic allopolyploid Cucumis hystrix × sativus has approximately the same
methylation density as the parents or the F1
hybrids.127
The direct involvement of DNA methylation in
expression changes between homeologs has also
been demonstrated. In the natural allotetraploid
A. suecica, the transcription factor TCP3 was
silenced when chemical inhibitors of DNA methyltransferases were applied,105 suggesting that methylation is important for proper TCP3 expression.
Another line of evidence comes from methyltransferase 1 (MET1) RNA interference (RNAi)
lines in the A. suecica background.128 Notably,
only 200 genes were found to be differentially expressed between A. suecica wild-type plants and
MET1 RNAi lines, and only 34 of these 200 genes
overlap with the 1400 genes with expression
changes in synthetic tetraploids between A. thaliana
and A. arenosa (the presumed diploid parents of
A. suecica).112 In addition, 33 of the 200 differentially
expressed genes in MET1 RNAi lines are pseudogenes or transposons.128 Thus, MET1-mediated
DNA methylation differences between diploids
and polyploids appear to be more relevant to
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Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 Moghe & Shiu
controlling heterochromatic states than to contributing to transcriptome shock. Nonetheless, the
DNA methylation machinery is complex,129 and
more loss-of-function studies will be necessary for
a more complete picture of the influence of DNA
methylation on transcriptome shock.
Similar to DNA methylation, there are indications that histone modifications likely play significant roles in transcriptome shock. Using diploids
derived from transgenic autotetraploid A. thaliana
containing epialleles that either silence or allow the
expression of a resistance gene marker,130 a screen
for mutants releasing the silencing effect of the epiallele resulted in identification of loss-of-function
alleles in DECREASE IN DNA METHYLATION1
(DDM1) and HOMOLOGOUS GENE SILENCING1
(HOG1).131 Global changes in both DNA and histone methylation in the mutant background likely
contribute to silencing release. Although the epiallele action appears to be tied to the ploidy level,130
it remains to be determined whether the mutations in DDM1 and/or HOG1 contribute to similar phenomenon in a polyploidy background. In
addition to histone methylation, histone acetylation is implicated in transcriptome shock based on
studies of a histone deacetylase (ATHD1/ATHDA19)
mutant.132 Genes differentially regulated between
wild-type and ATHD1/ATHDA19 mutants overlap
significantly with genes differentially expressed between synthetic allopolyploid Arabidopsis and the
diploid parents.
The expression profiles of small RNAs, including
both micro-RNA and small interfering (both repeatassociated and trans-acting) RNAs, are affected
postpolyploidization (Fig. 3C). Such changes have
been documented in resynthesized allotetraploids
of A. thaliana × arenosa133 and Ae. tauschii ×
Triticum turgidum.134 As with DNA methylation
and histone modification, mutants in genes controlling small RNA biogenesis have been used to
study the impact of small RNA on transcription
and other phenotypic changes associated with polyploidy. RNAi lines interfering with the expression of
DICER LIKE-1 and ARGONAUTE 1, both involved
in small RNA biogenesis, have been created.135 Similar to the DNA methylation machinery, the complexity of epigenetic regulatory machinery in plants
is well appreciated.136 Additional loss-of-function
studies in polyploids will be highly informative.
In addition to their effect on transcription, as dis-
Plant polyploidy
cussed earlier, small RNAs may contribute to the
phenomenon of genome dominance.66 Aside from
transcriptional regulatory mechanisms, a recent
study examining ribosome-associated transcripts in
the recently formed (100,000 years ago) allotetraploid Glycine dolichocarpa indicated that transcripts subject to translational regulation tend to
be retained homeologs from an ancient wholegenome duplication event.137 It will be interesting to perform similar experiments in the diploid
parents and F1s to assess the impact of genome
doubling and genome merger on translational
regulation.
Perturbations of biological networks due to
polyploidy
Preferential retention and loss of genes, coupled
with extensive transcriptional and genomic changes
postpolyploidy, necessarily leads to alterations in
the molecular networks operating in the organism (Fig. 4). The network-level changes, in turn,
are expected to affect the phenotype of the organism. Such phenotypic impact has been reviewed
recently, using flowering time as an example.120
While most retention/loss events may be random,
there is evidence for preferential retention/loss of
certain types of genes. According to the gene balance hypothesis, the random loss of genes can result in a perturbation of stoichiometric relationships between gene products, leading to genomic
imbalance in a newly created polyploid and loss of
fitness.138 Hence, neopolyploid lineages that retain
certain types of genes may be able to better establish
themselves.
There is a growing body of literature discussing
the functional evolution of gene duplicates derived
from whole-genome duplication.76,138,139 For example, studies in B. rapa show that genes involved in circadian rhythm such as CIRCADIAN CLOCK ASSOCIATED 1 and LATE ELONGATED HYPOCOTYL
were preferentially retained postpolyploidization,
leading to an altered flowering time pathway.140
Such preferential retention was also found in
A. thaliana genes duplicated in the ␣ polyploidization event 50–65 Ma related to specific metabolic
pathways,141 as well as MADS-box genes involved in
various aspects of plant development.142 In poplar,
two duplicate genes, FLOWERING LOCUS T1 and
T2, derived from a paleopolyploidization event
are expressed at different times of the year and
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Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 9
Plant polyploidy
Moghe & Shiu
Interacting
partners
Genetic/physical
interaction or
co-expression
found that, despite 100 Myr of evolution, genetic
redundancy may still exist between several pairs of
duplicated genes, although there is evidence of significant partitioning of ancestral functions and gain
of new functions between these pairs.147
Homeologous
genes
Gene 1
P1
Impact of polyploidy on morphology and
physiology
P2
Homeologous
genes
Gene 2
Polyploidization
Divergence
Sub-functionalization
Neo-functionalization
Gene loss
Figure 4. Effect of polyploidy on gene networks. P1, parent 1;
P2, parent 2. An example of two genes—gene 1 and gene 2—in
parents P1 and P2 is shown. Each square represents a genetic or
physical interaction of gene 1 or gene 2. After polyploidization,
over the course of several generations, the network topology
begins to evolve. If both of the homeologous copies do not undergo gene loss through deletion or pseudo-genization and are
retained, divergence between them will lead to eventual subfunctionalization or neofunctionalization between duplicates,
changing the network topology and, possibly, network function.
have distinct roles in influencing flowering time
and vegetative growth.143 Subfunctionalization of
two polyploidization-derived phytochrome (PHY)
genes in maize—PHY B1 and B2—where the two
proteins have both overlapping and unique functions in seedlings and adult plants, is also known.144
Most of our understanding of the impact of
polyploidy on biological network evolution comes
from budding yeast (Saccharomyces cerevisiae) that
underwent a polyploidization event 100 Ma.145
This polyploidization event was associated with
a large-scale rewiring of the transcriptional network through changes in cis-regulatory motif usage,
which led to the evolution of faculatively anaerobic
postpolyploidization species, whereas several prepolyploidization species are aerobic.146 It was also
10
Morphological alterations
Some fundamental commonalities in morphology
and physiology are found among polyploid species;
however, the specific outcomes of a particular polyploidization event can vary widely between taxa (reviewed in Ref. 148). Anatomically, polyploids have
larger cell volumes, stomatal guard cells, pollen, and
seeds compared to diploids. Polyploids also have
broader and thicker leaves with fewer stems per plant
(reviewed in Ref. 149). Such changes may affect processes such as water relations, gas exchange, cold
tolerance, and shade tolerance of polyploid plants.1
Some phenotypic modifications may occur owing
to increased DNA content in the nucleus and a need
to maintain a particular nuclear-to-cytoplasmic volume ratio, while other changes may be induced by
genome restructuring and regulatory and networklevel modifications. For example, in a synthetic autotetraploid line of A. thaliana, larger aerial organs
compared to the diploid are the result of faster expansion rates and a longer expansion duration during cell division. Such behavior is due to upregulation of cyclin-dependent kinases in tetraploids
compared to diploids.150
Polyploidization is also expected to affect the
morphology of the reproductive system by affecting the size of the flower, the relative sizes of petals,
the spatial relationships between different floral organs, and the flowering time of the plant.151 These
changes can influence pollinator preferences. In addition to morphological changes in reproductive
structure, polyploidy may lead to the breakdown
of self-incompatibility. In the Solanaceae family,
polyploid species are significantly more likely to be
self-compatible than diploid species.152 A broader
study of 235 angiosperm species also found a significant association between polyploidy and selfing.153
However, a study on self-sterility data from 1266
angiosperm species contrasting self-compatible,
self-incompatible, and mix-mating groups found
that polyploid species did not tend to be more
self-compatible compared to diploids.154 If the
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Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 Moghe & Shiu
results of the latter study are true, there may be
a short-term breakdown of self-incompatability
in the neopolyploid, enabling it to establish its
population,154 after which self-compatibility is
reestablished. Alternatively, other features associated with the mating system such as inflorescence
size, floral display, and pollinator behavior may
minimize the effect of reproductive isolation due
to polyploidization in neopolyploids. Nonetheless,
one issue with the 1266-species study is that phylogenetic relationships are not considered,154 although there is nonrandom association between
phylogeny and mating system. In addition, the selfing rate was binned into three broad categories.
On the other hand, the 235-species study considered species phylogeny and modeled selfing rate as a
continuous variable despite a smaller sample size.153
Thus, the lack of correlation between polyploidy and
selfing rate in the 1266-species study can be due to
confounding factors of phylogenetic relations and
selfing-rate binning.
Physiological changes and stress tolerance
Polyploidy significantly influences photosynthesis
(reviewed in Ref. 155), and this effect is particularly
obvious under stress conditions. In greenhousegrown Betula papyrifer, water stress treatment leads
to a complete cessation of photosynthesis in diploids
but not in penta- and hexapolyploids. Such behavior can be partly attributed to earlier stomata closure in diploids.156 However, it remains unclear if
the photosynthetic activity under water deficit in
polyploids is advantageous, because it would contribute to carbon fixation even under stress, or
whether it is detrimental due to water loss. Under
excess light, the capacity for photoprotection and
nonphotosynthetic electron transport are higher in
the natural allotetraploid G. dolichocarpa than in its
diploid progenitors.157 This enhanced photoprotection appears beneficial because the allotetraploid is
reported to photobleach later than its diploid progenitors.
Because of the prevalence of polyploid plants
and the perceived broader ecological tolerance (reviewed in Ref. 149), one potential physiological consequence of polyploidy is its increased tolerance to
environmental stress. However, the degree of environmental stress tolerance does not necessarily correlate with cytotypes (see Ref. 158, and references
therein). Taking drought tolerance as an example,
Plant polyploidy
in a comparison between the fireweed Chamerion
angustifolium diploids and tetraploids (both natural and synthetic) in a controlled environment, the
natural tetraploids took 20–30% longer to wilt compared to both diploid and synthetic tetraploids,158
consistent with the higher xylem hydraulic conductivity observed in natural tetraploids. However, the
vulnerability of stems to drought-induced cavitation was similar among C. angustifolium cytotypes.
In field grown tetraploid and hexaploid Atriplex
canescens,159 the leaf-specific hydraulic conductivity as well as susceptibility to cavitation was lower
in plants with higher ploidy levels. The inconsistency between earlier studies and the two recent ones
highlighted above can be attributed to, for example,
differences in whether the studies were conducted
in controlled or natural environments, how physiological measurements were taken, and how and
when the polyploids were established.
Polyploids are also hypothesized to be more resistant to pathogens.160 Mathematical models of interactions between pathogens and either diploids or
neopolyploids have shown that newly formed polyploid populations of hosts are expected to be more
resistant.161 However, similar to studies of the relationships between polyploidy and abiotic stress
tolerance, the few empirical studies conducted so
far have generated mixed results.162,163 For plant–
insect interactions, the autotetraploid gooseberryleaf alumroot Heuchera grossulariifolia is more likely
to be attacked by the specialist moth herbivore Greya
politella than the diploid.164 In another field study,
the resistance of H. grossulariifolia to three moth
species (G. politella, G. piperella, and Eupithecia misturata) was tested.165 Interestingly, G. piperella tends
to attack and lay eggs on diploids, suggesting herbivore species may provide selective pressure differently due to differences in ploidy levels. Nonetheless,
it remains unclear whether there are additional genetic differences independent from polyploidy that
contribute to the difference.
Polyploidy and adaptation
Survival in adverse environments
The question of whether polyploidization contributes to the long-term evolutionary success of
plant species has been raised repeatedly since
Stebbins.166 There are already a number of excellent reviews on this topic;17,149,167,168 thus, in this
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Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 11
Plant polyploidy
Moghe & Shiu
section, we focus on providing a summary of earlier
findings and a discussion of some recent results.
Polyploids are more frequent at higher elevations and higher latitudes and may be more tolerant
to dry conditions,38,158,169 suggesting a fitness advantage for polyploids in those environments. The
timing of multiple polyploidization events in angiosperms coincides with the timing of the creation
of the Cretaceous–Tertiary boundary.170 This coincidence has led to the hypothesis that species with
genome doubling could adapt better to the changing environment than their diploid relatives during
the mass extinction event. However, considering the
influence of environmental factors on unreduced
gamete formation, it is also possible that the intense
climatic changes during the mass extinction event
may have increased the frequency of unreduced gamete formation, creating polyploids at a faster rate
than normal. In addition, polyploids were found
to be more successful in colonizing the Arctic after deglaciation than diploids.169 A study sampling
640 endangered and 81 invasive species worldwide
has led to the conclusion that endangered species
tend to be diploids while invasive species tend to be
polyploids, suggesting that polyploidization may increase tolerance to diverse ecological conditions.171
It has also been shown that polyploid A. thaliana
accessions are more tolerant to and have better reproductive success under high salinity compared to
diploid cytotypes.172 It will be particularly interesting to determine if such high-salinity adaptation can
also be observed under field conditions.
Not all findings support the notion that polyploids tend to survive better in adverse environments. A 1940 study of 100 polyploid and diploid
plant species found no correlation between polyploidy and winter hardiness.173 More recently, in a
study of two diploid and one polyploid species from
each of 144 North American plant genera, no association was found between ploidy level and species
range area, minimum/maximum temperature, precipitation, or latitudes.174 Furthermore, as discussed
in the previous section, physiological changes associated with polyploidy do not necessarily confer
superior tolerance and/or resistance to abiotic and
biotic factors. These observations suggest that the
relationship between polyploidy and adaptation is
quite complex and may depend on not only the
species undergoing polyploidization but also on the
environment.
12
Local adaptation
In addition to the meta-analyses discussed earlier,
multiple studies provide specific examples of adaptation that are potentially attributable to polyploidy.
In H. grossulariifolia, pollinators visit tetraploid individuals more often than diploid ones.175 Similarly,
in the fireweed C. angustifolium, where populations
ranges of polyploid and diploid varieties overlap,
tetraploids have a disproportionately higher number of bee visits and a greater pollen-siring advantage compared to diploids.176,177 The invasiveness
of allotetraploid cordgrass S. anglica may indicate a
fitness increase due to polyploidy; however, this is
more likely a consequence of heterosis than genome
doubling.178
In wild yarrow Achillea borealis, which has
hexaploid and tetraploid populations occupying
nearby but different environments, there is clear
evidence of local adaptation, and polyploidization
is likely the initial trigger for diversification and
adaptation to a new habitat.179 Consistent with this
notion, the evolvability of the neo-autotetraploid
C. angustifolium is higher than both diploid and
established autotetraploids, suggesting that genome
doubling, without hybridization, may initially
alter evolutionary rate and contribute to adaptive
evolution.180 In addition, reciprocal transplant
experiments demonstrated that C. angustifolium
diploids and tetraploids survived best at their native
elevations.181 Overall, these observations suggest
that polyploidization may have positive fitness
consequences and can lead to adaptation to local or
regional environments.
Species richness
Polyploidy leads to instantaneous reproductive isolation of polyploid individuals through the phenomenon of minority cytotype exclusion,1,30 but
such individuals also possess a greater capacity for
functional innovation. If polyploids in general have
better fitness compared to diploids, the speciation rate of polyploids may be higher than that of
diploids. Multiple observations suggest a positive
association between species richness and percent
polyploid species in different plant clades.7,149,182
In addition, a large number of species in major
plant families, such as Poaceae,183 Asteraceae,184
Brassicaceae,57 and the subfamily Papilionoideae,185
have descended from a polyploid ancestor, suggesting a possible increase in diversification rates
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Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 Moghe & Shiu
postpolyploidization. Overall, it has been estimated
that a significant proportion, 15% of angiosperm
and 31% of fern speciation events, may have been
accompanied by ploidy increase.186 On the contrary,
some other studies provide conflicting results. One
study arrived at the conclusion that polyploids tend
to have lower diversification rates than diploids and
have a greater chance of extinction.187 It has also
been estimated that only 2–4% of polyploidization events in angiosperms have actually resulted
in speciation.149 In addition, analyses of genome
sequences of flowering plants suggests that the difference in the estimated timing between successive
detectable paleopolyploidization events is 10–30
Myr in flowering plant lineages.5 Thus, considering the abundance of polyploids among flowering
plants, most of the polyploidization events were not
recorded in the genomes of extant species. The inference is that most of these polyploids have gone
extinct.
These observations suggest two possibilities—
either a majority of the polyploid lineages indeed
go extinct, making them evolutionary dead ends,166
or they hybridize with their parental species, creating lineages of mixed ploidy. It seems that both
of these scenarios may occur in nature. Hybridization between polyploids and their diploid ancestors
has been reported in natural populations of multiple species such as C. angustifolium,188 species of
the genera Epidendrum189 and Jacobaea,190 and several others,191 in regions known as hybrid zones.191
Mathematical simulations on whether a triploid hybrid between an autotetraploid and a diploid can
help a tetraploid population to establish suggest
that even partially fit triploids can assist in longterm tetraploid fixation.19 Thus, polyploidization
may not necessarily be an evolutionary dead end
but may create interesting possibilities for further
innovation in the lineage.
Conclusions
In this article, we highlight the findings that the formation of a polyploid is associated with extensive
changes at the genomic, epigenetic, transcriptional,
and network levels. These genomic, transcriptomic,
and other omic changes must have contributed to
morphological, physiological, and ecological phenotypic differences between polyploids and their
diploid progenitors. However, the exact molecular changes responsible for the phenotypic differ-
Plant polyploidy
ences between cytotypes remain unclear in most
cases. Also, we have a relatively better understanding of the molecular and phenotypic consequences
of allopolyploidy than autopolyploidy, and studies
comparing and contrasting mechanisms of molecular evolution in these two forms of polyploidy are
lacking. For example, it is not clear whether the
rate of neo- and subfunctionalization and pseudogenization differ between auto- and allopolyploids,
given the homeologs in allopolyploids are already
slightly divergent from each other. Also, given that
the extent of functional redundancy is higher in autopolyploids, do mutations have a stronger deleterious effects in allopolyploids than autopolyploids?
Additional research would be needed to address
these questions.
In a review by Soltis et al.,13 a number of intriguing questions are raised regarding what we still
do not know about polyploidy. To expand on the
long list, one challenge lies in establishing the genetic basis in cases where polyploids are shown to
be successful. Among the unknowns, one particularly challenging question is whether polyploids are
more successful than their diploid progenitors.13 A
related question concerns determining the ecological situations under which polyploidy is adaptive.
Theoretical considerations as well as empirical evidence have provided contradictory answers to these
two questions so far.17,149,166–168 Nevertheless, the
second question is more tractable, as it does not
require generalization and can be examined experimentally on a species-by-species basis. Polyploidy
is an extreme form of duplication and can be seen
as a mutation mechanism. Considering the nearly
neutral theory of molecular evolution,192 the null
hypothesis is that the effect of polyploidy is neutral
or nearly neutral. In this framework, we can test
under what situations (e.g., different abiotic/biotic
environments) the null hypothesis can be rejected.
Taking C. angustifolium as an example, diploid
and tetraploid varieties show significant differences
in drought tolerance.158 Given the understanding of
drought tolerance in model plants,193,194 a targeted
survey of candidate gene transcription in field conditions can potentially be informative. The optimal
timing for the assay is not trivial to determine, and
molecular changes other than transcription can be
more important. In addition, drought tolerance may
not be the main environmental factor, highlighting
the need to assess potentially relevant abiotic/biotic
C 2014 New York Academy of Sciences.
Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 13
Plant polyploidy
Moghe & Shiu
factors in controlled environments in addition to
the field. Nonetheless, a candidate gene approach
is a reasonable starting point. In case the candidate
gene approach does not bear fruit, because sampling
the omes—genomes, transcriptomes, proteomes, or
metabolomes—of nonmodel species is no longer
a rate-limiting step, a global study of molecular
changes may provide viable hypotheses for further testing. These considerations are not unique to
C. angustifolium but are relevant to other polyploid
study systems as well. We surmise that good experimental designs incorporating both molecular and
ecological considerations of polyploids have the potential to make the most impact in the near future.
Acknowledgments
We would like to thank Jonathan Wendel and Joshua
Udall for discussion, the two anonymous reviewers,
as well as the editor for providing critical feedback
on the manuscript. SHS is supported by National
Science Foundation Grants DEB-0919452 and IOS1126998.
Conflicts of interest
The authors declare no conflicts of interest.
References
1. Ramsey, J. & D.W. Schemske. 2002. Neopolyploidy in flowering plants. Annu. Rev. Ecol. Syst. 33: 589–639.
2. Tate, J.A., P.S. Soltis & D.E. Soltis. 2005. “Chapter 7: polyploidy in plants.” In The Evolution of the Genome. T. Ryan
Gregory, Ed.: 371–426. Academic Press.
3. Husband, B.C., S.J. Baldwin & J. Suda. 2013. “The incidence
of polyploidy in natural plant populations: major patterns
and evolutionary processes.” In Plant Genome Diversity.
Vol. 2. I.J. Leitch, J. Greilhuber, J. Dolezel, et al. Eds.: 255–
276. Vienna: Springer.
4. Friedman, W.E. 2009. The meaning of Darwin’s “abominable mystery.” Am. J. Bot. 96: 5–21.
5. Jiao, Y., N.J. Wickett, S. Ayyampalayam, et al. 2011. Ancestral polyploidy in seed plants and angiosperms. Nature
473: 97–100.
6. Ohno, S. 1970. Evolution by Gene Duplication. New York:
Springer-Verlag.
7. Soltis, D.E., V.A. Albert, J. Leebens-Mack, et al. 2009. Polyploidy and angiosperm diversification. Am. J. Bot. 96: 336–
348.
8. Gregory, R. & B. Mable. 2005. “Chapter 8—polyploidy in
animals”. In The Evolution of the Genome. Elsevier Academic Press.
9. Albertin, W. & P. Marullo. 2012. Polyploidy in fungi: evolution after whole-genome duplication. Proc. R. Soc. B 279:
2497–2509.
14
10. Soppa, J. 2013. Evolutionary advantages of polyploidy in
halophilic archaea. Biochem. Soc. Trans. 41: 339–343.
11. Innan, H. & F. Kondrashov. 2010. The evolution of gene duplications: classifying and distinguishing between models.
Nat. Rev. Genet. 11: 97–108.
12. Ramsey, J. & D.W. Schemske. 1998. Pathways, mechanisms,
and rates of polyploid formation in flowering plants. Annu.
Rev. Ecol. Syst. 29: 467–501.
13. Soltis, D.E., R.J.A. Buggs, J.J. Doyle & P.S. Soltis. 2010. What
we still don’t know about polyploidy. Taxon 59: 1387–1403.
14. Soltis, D.E. & P.S. Soltis. 2012. Polyploidy and Genome Evolution. New York: Springer.
15. Leitch, I.J., J. Greilhuber, J. Dolezel & J. Wendel. 2013. Plant
Genome Diversity. Vol. 2: Physical Structure, Behaviour and
Evolution of Plant Genomes. Vienna: Springer-Verlag.
16. Stöck, M. & D.K. Lamatsch. 2013. Trends in Polyploidy Research in Animals and Plants: Reprint of: Cytogenetic and
Genome Research 2013. Vol. 140, Issues 2–4. Basel: S. Karger.
17. Madlung, A. 2013. Polyploidy and its effect on evolutionary
success: old questions revisited with new tools. Heredity
110: 99–104.
18. Bretagnolle, F. & J.D. Thompson. 1995. Tansley Review
No. 78. Gametes with the stomatic chromosome number:
mechanisms of their formation and role in the evolution of
autopolypoid plants. New Phytol. 129: 1–22.
19. Husband, B.C. 2004. The role of triploid hybrids in the
evolutionary dynamics of mixed-ploidy populations. Biol.
J. Linn. Soc. 82: 537–546.
20. Yamauchi, A., A. Hosokawa, H. Nagata & M. Shimoda.
2004. Triploid bridge and role of parthenogenesis in the
evolution of autopolyploidy. Am. Nat. 164: 101–112.
21. Suda, J. & T. Herben. 2013. Ploidy frequencies in plants
with ploidy heterogeneity: fitting a general gametic model
to empirical population data. Proc. R. Soc. B 280: 20122387.
22. Fox, D.T. & R.J. Duronio. 2013. Endoreplication and polyploidy: insights into development and disease. Development
140: 3–12.
23. Brownfield, L. & C. Köhler. 2011. Unreduced gamete formation in plants: mechanisms and prospects. J. Exp. Bot.
62: 1659–1668.
24. De Storme, N. & D. Geelen. 2013. Sexual polyploidization
in plants–cytological mechanisms and molecular regulation. New Phytol. 198: 670–684.
25. Ravi, M., M.P.A. Marimuthu & I. Siddiqi. 2008. Gamete formation without meiosis in Arabidopsis. Nature 451: 1121–
1124.
26. Siddiqi, I., G. Ganesh, U. Grossniklaus & V. Subbiah. 2000.
The dyad gene is required for progression through female
meiosis in Arabidopsis. Development 127: 197–207.
27. Mercier, R., D. Vezon, E. Bullier, et al. 2001. SWITCH1
(SWI1): a novel protein required for the establishment of
sister chromatid cohesion and for bivalent formation at
meiosis. Genes Dev. 15: 1859–1871.
28. De Storme, N., G.P. Copenhaver & D. Geelen. 2012. Production of diploid male gametes in Arabidopsis by coldinduced destabilization of post-meiotic radial microtubule
arrays. Plant Physiol. 160: 1808–1826.
29. Marks, G.E. 1966. The origin and significance of intraspecific polyploidy: experimental evidence from Solanum chacoense. Evolution 20: 552–557.
C 2014 New York Academy of Sciences.
Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 Moghe & Shiu
30. Levin, D.A. 1975. Minority cytotype exclusion in local plant
populations. Taxon 24: 35–43.
31. Kradolfer, D., P. Wolff, H. Jiang, et al. 2013. An imprinted
gene underlies postzygotic reproductive isolation in Arabidopsis thaliana. Dev. Cell 26: 525–535.
32. Parrott, W.A. & R.R. Smith. 1986. Recurrent selection for
2n pollen formation in red clover. Crop Sci. 26: 1132–1135.
33. Tavoletti, S., A. Mariani & F. Veronesi. 1991. Phenotypic
recurrent selection for 2n pollen and 2n egg production in
diploid alfalfa. Euphytica 57: 97–102.
34. Tischler, G. 1935. Die Bedeutung der Polyploidie für die
Verbreitung der Angiospermen. Bot. Jahrbucher 76: 1–36.
35. Sakamura, T. 1920. Experimentelle studien über die Zell
und Kerntcilung, mit besonderer Rücksicht auf Form,
Grösse und Zahl dei Chromosomen. J. Coll. Sci. Imp. Univ.
Tokyo 39: 221.
36. Randolph, L.F. 1932. Some effects of high temperature on
polyploidy and other variations in maize. Proc. Natl. Acad.
Sci. U. S. A. 18: 222–229.
37. Pécrix, Y., G. Rallo, H. Folzer, et al. 2011. Polyploidization
mechanisms: temperature environment can induce diploid
gamete formation in Rosa sp. J. Exp. Bot. 62: 3587–3597.
38. Grant, V. 1981. Plant Speciation. New York: Columbia University Press.
39. Madlung, A., A.P. Tyagi, B. Watson, et al. 2005. Genomic
changes in synthetic Arabidopsis polyploids. Plant J. 41:
221–230.
40. Gaeta, R.T., J.C. Pires, F. Iniguez-Luy, et al. 2007. Genomic
changes in resynthesized Brassica napus and their effect on
gene expression and phenotype. Plant Cell 19: 3403–3417.
41. Buggs, R.J.A., A.N. Doust, J.A. Tate, et al. 2009. Gene loss
and silencing in Tragopogon miscellus (Asteraceae): comparison of natural and synthetic allotetraploids. Heredity
103: 73–81.
42. Xiong, Z., R.T. Gaeta & J.C. Pires. 2011. Homoeologous shuffling and chromosome compensation maintain
genome balance in resynthesized allopolyploid Brassica napus. Proc. Natl. Acad. Sci. U. S. A. 108: 7908–7913.
43. Chester, M., J.P. Gallagher, V.V. Symonds, et al. 2012. Extensive chromosomal variation in a recently formed natural
allopolyploid species, Tragopogon miscellus (Asteraceae).
Proc. Natl. Acad. Sci. U. S. A. 109: 1176–1181.
44. Mestiri, I., V. Chagué, A.-M. Tanguy, et al. 2010. Newly
synthesized wheat allohexaploids display progenitordependent meiotic stability and aneuploidy but structural
genomic additivity. New Phytol. 186: 86–101.
45. Matsushita, S.C., A.P. Tyagi, G.M. Thornton, et al. 2012.
Allopolyploidization lays the foundation for evolution of
distinct populations: evidence from analysis of synthetic
Arabidopsis allohexaploids. Genetics 191: 535–547.
46. Lamm, R. 1945. Cytogenetic studies in Solanum, sect. Tuberarium. Hereditas 31: 1–129.
47. Shaver, D.L. 1962. A study of aneuploidy in autotetraploid
maize. Can. J. Genet. Cytol. 4: 226–233.
48. Henry, I.M., B.P. Dilkes, A. Tyagi, et al. 2014. The BOY
NAMED SUE quantitative trait locus confers increased
meiotic stability to an adapted natural allopolyploid of Arabidopsis. Plant Cell 26: 181–194.
Plant polyploidy
49. Martelotto, L.G., J.P.A. Ortiz, J. Stein, et al. 2007. Genome
rearrangements derived from autopolyploidization in Paspalum sp. Plant Sci. 172: 970–977.
50. Eilam, T., Y. Anikster, E. Millet, et al. 2009. Genome size in
natural and synthetic autopolyploids and in a natural segmental allopolyploid of several Triticeae species. Genome
52: 275–285.
51. Ozkan, H., M. Tuna & D.W. Galbraith. 2006. No DNA loss
in autotetraploids of Arabidopsis thaliana. Plant Breed. 125:
288–291.
52. Li, W.H., T. Gojobori & M. Nei. 1981. Pseudogenes as a
paradigm of neutral evolution. Nature 292: 237–239.
53. Blanc, G., K. Hokamp & K.H. Wolfe. 2003. A recent polyploidy superimposed on older large-scale duplications in
the Arabidopsis genome. Genome Res. 13: 137–144.
54. Schmutz, J., S.B. Cannon, J. Schlueter, et al. 2010. Genome
sequence of the palaeopolyploid soybean. Nature 463: 178–
183.
55. Yang, Y.W., K.N. Lai, P.Y. Tai & W.H. Li. 1999. Rates of
nucleotide substitution in angiosperm mitochondrial DNA
sequences and dates of divergence between Brassica and
other angiosperm lineages. J. Mol. Evol. 48: 597–604.
56. Town, C.D., F. Cheung, R. Maiti, et al. 2006. Comparative
genomics of Brassica oleracea and Arabidopsis thaliana
reveal gene loss, fragmentation, and dispersal after polyploidy. Plant Cell 18: 1348–1359.
57. Beilstein, M.A., N.S. Nagalingum, M.D. Clements, et al.
2010. Dated molecular phylogenies indicate a Miocene origin for Arabidopsis thaliana. Proc. Natl. Acad. Sci. U. S. A.
107: 18724–18728.
58. Wang, X., H. Wang, J. Wang, et al. 2011. The genome of the
mesopolyploid crop species Brassica rapa. Nat. Genet. 43:
1035–1039.
59. Moghe, G.D., D.E. Hufnagel, H. Tang, et al. In press. Consequences of whole genome triplication as revealed by comparative genomic analyses of the wild radish Raphanus
raphanistrum and three other Brassicaceae species. Plant
Cell.
60. Langham, R.J., J. Walsh, M. Dunn, et al. 2004. Genomic duplication, fractionation and the origin of regulatory novelty.
Genetics 166: 935–945.
61. Freeling, M. 2009. Bias in plant gene content following
different sorts of duplication: tandem, whole-genome, segmental, or by transposition. Annu. Rev. Plant Biol. 60: 433–
453.
62. Thomas, B.C., B. Pedersen & M. Freeling. 2006. Following
tetraploidy in an Arabidopsis ancestor, genes were removed
preferentially from one homeolog leaving clusters enriched
in dose-sensitive genes. Genome Res. 16: 934–946.
63. Schnable, J.C., N.M. Springer & M. Freeling. 2011. Differentiation of the maize subgenomes by genome dominance
and both ancient and ongoing gene loss. Proc. Natl. Acad.
Sci. U. S. A. 108: 4069–4074.
64. Cheng, F., J. Wu, L. Fang, et al. 2012. Biased gene fractionation and dominant gene expression among the subgenomes
of Brassica rapa. PLoS One 7: e36442.
65. Tang, H., M.R. Woodhouse, F. Cheng, et al. 2012. Altered
patterns of fractionation and exon deletions in Brassica rapa
C 2014 New York Academy of Sciences.
Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 15
Plant polyploidy
66.
67.
68.
69.
70.
71.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
82.
16
Moghe & Shiu
support a two-step model of paleohexaploidy. Genetics 190:
1563–1574.
Woodhouse, M.R., F. Cheng, J.C. Pires, et al. 2014. Origin,
inheritance, and gene regulatory consequences of genome
dominance in polyploids. Proc. Natl. Acad. Sci. U. S. A. 111:
5283–5288.
Blanc, G. & K.H. Wolfe. 2004. Functional divergence of
duplicated genes formed by polyploidy during Arabidopsis
evolution. Plant Cell 16: 1679–1691.
Hanada, K., C. Zou, M.D. Lehti-Shiu, et al. 2008. Importance of lineage-specific expansion of plant tandem duplicates in the adaptive response to environmental stimuli.
Plant Physiol. 148: 993–1003.
Chapman, B.A., J.E. Bowers, F.A. Feltus & A.H. Paterson.
2006. Buffering of crucial functions by paleologous duplicated genes may contribute cyclicality to angiosperm
genome duplication. Proc. Natl. Acad. Sci. U. S. A. 103:
2730–2735.
Jiang, W.K., Y.L. Liu, E.H. Xia & L.Z. Gao. 2013. Prevalent
role of gene features in determining evolutionary fates of
whole-genome duplication duplicated genes in flowering
plants. Plant Physiol. 161: 1844–1861.
Pál, C., B. Papp & L.D. Hurst. 2001. Highly expressed genes
in yeast evolve slowly. Genetics 158: 927–931.
Chang, P.L., B.P. Dilkes, M. McMahon, et al. 2010.
Homoeolog-specific retention and use in allotetraploid
Arabidopsis suecica depends on parent of origin and network partners. Genome Biol. 11: R125.
Force, A., M. Lynch, F.B. Pickett, et al. 1999. Preservation
of duplicate genes by complementary, degenerative mutations. Genetics 151: 1531–1545.
Freeling, M. & B.C. Thomas. 2006. Gene-balanced duplications, like tetraploidy, provide predictable drive to increase
morphological complexity. Genome Res. 16: 805–814.
Birchler, J.A. & R.A. Veitia. 2007. The gene balance hypothesis: from classical genetics to modern genomics. Plant Cell
19: 395–402.
Conant, G.C. & K.H. Wolfe. 2008. Turning a hobby into
a job: how duplicated genes find new functions. Nat. Rev.
Genet. 9: 938–950.
Des Marais, D.L. & M.D. Rausher. 2008. Escape from adaptive conflict after duplication in an anthocyanin pathway
gene. Nature 454: 762–765.
Edger, P.P. & J.C. Pires. 2009. Gene and genome duplications: the impact of dosage-sensitivity on the fate of nuclear
genes. Chromosome Res. 17: 699–717.
Kondrashov, F.A. 2012. Gene duplication as a mechanism
of genomic adaptation to a changing environment. Proc. R.
Soc. B 279: 5048–5057.
Haldane, J.B.S. 1932. The Causes of Evolution. Princeton
University Press.
Tsai, H., V. Missirian, K.J. Ngo, et al. 2013. Production
of a high-efficiency TILLING population through polyploidization. Plant Physiol. 161: 1604–1614.
Akhunov, E.D., S. Sehgal, H. Liang, et al. 2013. Comparative
analysis of syntenic genes in grass genomes reveals accelerated rates of gene structure and coding sequence evolution
in polyploid wheat. Plant Physiol. 161: 252–265.
83. Parisod, C., K. Alix, J. Just, et al. 2010. Impact of transposable elements on the organization and function of allopolyploid genomes. New Phytol. 186: 37–45.
84. Parisod, C. & N. Senerchia. 2012. “Responses of transposable elements to polyploidy.” In Plant Transposable Elements. M.A. Grandbastien & J.M. Casacuberta, Eds.: 147–
168. Berlin, Heidelberg: Springer.
85. Ben-David, S., B. Yaakov & K. Kashkush. 2013. Genomewide analysis of short interspersed nuclear elements SINES
revealed high sequence conservation, gene association and
retrotranspositional activity in wheat. Plant J. 76: 201–210.
86. Hazzouri, K.M., A. Mohajer, S.I. Dejak, et al. 2008. Contrasting patterns of transposable-element insertion polymorphism and nucleotide diversity in autotetraploid and
allotetraploid Arabidopsis species. Genetics 179: 581–592.
87. Beaulieu, J., M. Jean & F. Belzile. 2009. The allotetraploid
Arabidopsis thaliana-Arabidopsis lyrata subsp. petraea as an
alternative model system for the study of polyploidy in
plants. Mol. Genet. Genomics 281: 421–435.
88. Parisod, C., A. Salmon, T. Zerjal, et al. 2009. Rapid structural and epigenetic reorganization near transposable elements in hybrid and allopolyploid genomes in Spartina.
New Phytol. 184: 1003–1015.
89. Petit, M., C. Guidat, J. Daniel, et al. 2010. Mobilization of
retrotransposons in synthetic allotetraploid tobacco. New
Phytol. 186: 135–147.
90. Chantret, N., J. Salse, F. Sabot, et al. 2005. Molecular basis of evolutionary events that shaped the hardness locus in diploid and polyploid wheat species (Triticum and
Aegilops). Plant Cell 17: 1033–1045.
91. Bruggmann, R., A.K. Bharti, H. Gundlach, et al. 2006. Uneven chromosome contraction and expansion in the maize
genome. Genome Res. 16: 1241–1251.
92. Adams, K.L. & J.F. Wendel. 2005. Novel patterns of gene
expression in polyploid plants. Trends Genet. 21: 539–543.
93. Adams, K.L. 2007. Evolution of duplicate gene expression
in polyploid and hybrid plants. J. Hered. 98: 136–141.
94. Jackson, S. & Z.J. Chen. 2010. Genomic and expression
plasticity of polyploidy. Curr. Opin. Plant Biol. 13: 153–
159.
95. Birchler, J.A. 2012. Insights from paleogenomic and population studies into the consequences of dosage sensitive
gene expression in plants. Curr. Opin. Plant Biol. 15: 544–
548.
96. Hegarty, M., J. Coate, S. Sherman-Broyles, et al. 2013.
Lessons from natural and artificial polyploids in higher
plants. Cytogenet. Genome Res. 140: 204–225.
97. Zou, C., M.D. Lehti-Shiu, M. Thomashow & S.-H. Shiu.
2009. Evolution of stress-regulated gene expression in
duplicate genes of Arabidopsis thaliana. PLoS Genet. 5:
e1000581.
98. Hegarty, M.J., G.L. Barker, I.D. Wilson, et al. 2006. Transcriptome shock after interspecific hybridization in Senecio
is ameliorated by genome duplication. Curr. Biol. 16: 1652–
1659.
99. Wang, J., L. Tian, H.S. Lee, et al. 2006. Genomewide nonadditive gene regulation in Arabidopsis allotetraploids. Genetics 172: 507–517.
C 2014 New York Academy of Sciences.
Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 Moghe & Shiu
100. Xu, Y., L. Zhong, X. Wu, et al. 2009. Rapid alterations of gene
expression and cytosine methylation in newly synthesized
Brassica napus allopolyploids. Planta 229: 471–483.
101. Flagel, L., J. Udall, D. Nettleton & J. Wendel. 2008. Duplicate
gene expression in allopolyploid Gossypium reveals two
temporally distinct phases of expression evolution. BMC
Biol. 6: 16.
102. Chelaifa, H., V. Chagué, S. Chalabi, et al. 2013. Prevalence
of gene expression additivity in genetically stable wheat
allohexaploids. New Phytol. 197: 730–736.
103. Gianinetti, A. 2013. A criticism of the value of midparent
in polyploidization. J. Exp. Bot. 64: 4119–4129.
104. Grover, C.E., J.P. Gallagher, E.P. Szadkowski, et al. 2012. Homoeolog expression bias and expression level dominance
in allopolyploids. New Phytol. 196: 966–971.
105. Lee, H.S. & Z.J. Chen. 2001. Protein-coding genes are epigenetically regulated in Arabidopsis polyploids. Proc. Natl.
Acad. Sci. U. S. A. 98: 6753–6758.
106. Yoo, M.-J., E. Szadkowski & J.F. Wendel. 2013. Homoeolog expression bias and expression level dominance in
allopolyploid cotton. Heredity 110: 171–180.
107. Wendel, J.F. & R.C. Cronn. 2003. “Polyploidy and the evolutionary history of cotton.” In Advances in Agronomy. pp.
139–186. Academic Press.
108. Rapp, R.A., J.A. Udall & J.F. Wendel. 2009. Genomic expression dominance in allopolyploids. BMC Biol. 7: 18.
109. Xu, C., Y. Bai, X. Lin, et al. 2014. Genome-wide disruption
of gene expression in allopolyploids but not hybrids of rice
subspecies. Mol. Biol. Evol.
110. Ownbey, M. 1950. Natural hybridization and amphiploidy
in the genus Tragopogon. Am. J. Bot. 37: 487–499.
111. Lowe, A. & R.J. Abbott. 2003. A new British species Senecio
eboracensis (Asteraceae) another hybrid derivative of S.
vulgaris L. and S. squalidus L. Watsonia 24: 375–388.
112. Wang, J., L. Tian, H.S. Lee, et al. 2006. Genomewide nonadditive gene regulation in Arabidopsis allotetraploids. Genetics 172: 507–517.
113. Church, S.A. & E.J. Spaulding. 2009. Gene expression in
a wild autopolyploid sunflower series. J. Hered. 100: 491–
495.
114. Chelaifa, H., A. Monnier & M. Ainouche. 2010. Transcriptomic changes following recent natural hybridization
and allopolyploidy in the salt marsh species Spartina ×
townsendii and Spartina anglica (Poaceae). New Phytol. 186:
161–174.
115. Yang, C., Y. Kawahara, H. Mizuno, et al. 2012. Independent
domestication of Asian rice followed by gene flow from
japonica to indica. Mol. Biol. Evol. 29: 1471–1479.
116. Chen, Z.J. 2007. Genetic and epigenetic mechanisms for
gene expression and phenotypic variation in plant polyploids. Annu. Rev. Plant Biol. 58: 377–406.
117. Comai, L. 2000. Genetic and epigenetic interactions in allopolyploid plants. Plant Mol. Biol. 43: 387–399.
118. Doyle, J.J., L.E. Flagel, A.H. Paterson, et al. 2008. Evolutionary genetics of genome merger and doubling in plants.
Annu. Rev. Genet. 42: 443–461.
119. Chen, Z.J. 2010. Molecular mechanisms of polyploidy and
hybrid vigor. Trends Plant Sci. 15: 57.
Plant polyploidy
120. Mayfield, D., Z.J. Chen & J.C. Pires. 2011. Epigenetic regulation of flowering time in polyploids. Curr. Opin. Plant
Biol. 14: 174–178.
121. Ng, D.W., J. Lu & Z.J. Chen. 2012. Big roles for small RNAs
in polyploidy, hybrid vigor, and hybrid incompatibility.
Curr. Opin. Plant Biol. 15: 154–161.
122. Madlung, A. & J.F. Wendel. 2013. Genetic and epigenetic
aspects of polyploid evolution in plants. Cytogenet. Genome
Res. 140: 270–285.
123. Lukens, L.N., J.C. Pires, E. Leon, et al. 2006. Patterns of
sequence loss and cytosine methylation within a population
of newly resynthesized Brassica napus allopolyploids. Plant
Physiol. 140: 336–348.
124. Shaked, H., K. Kashkush, H. Ozkan, et al. 2001. Sequence
elimination and cytosine methylation are rapid and reproducible responses of the genome to wide hybridization and
allopolyploidy in wheat. Plant Cell 13: 1749–1760.
125. Salmon, A., M.L. Ainouche & J.F. Wendel. 2005. Genetic
and epigenetic consequences of recent hybridization and
polyploidy in Spartina (Poaceae). Mol. Ecol. 14: 1163–1175.
126. Madlung, A., R.W. Masuelli, B. Watson, et al. 2002. Remodeling of DNA methylation and phenotypic and transcriptional changes in synthetic Arabidopsis allotetraploids.
Plant Physiol. 129: 733–746.
127. Chen, L. & J. Chen. 2008. Changes of cytosine methylation induced by wide hybridization and allopolyploidy in
Cucumis. Genome 51: 789–799.
128. Chen, M., M. Ha, E. Lackey, et al. 2008. RNAi of met1
reduces DNA methylation and induces genome-specific
changes in gene expression and centromeric small RNA
accumulation in Arabidopsis allopolyploids. Genetics 178:
1845–1858.
129. Law, J.A. & S.E. Jacobsen. 2010. Establishing, maintaining
and modifying DNA methylation patterns in plants and
animals. Nat. Rev. Genet. 11: 204–220.
130. Mittelsten Scheid, O., K. Afsar & J. Paszkowski. 2003. Formation of stable epialleles and their paramutation-like interaction in tetraploid Arabidopsis thaliana. Nat. Genet. 34:
450–454.
131. Baubec, T., H.Q. Dinh, A. Pecinka, et al. 2010. Cooperation
of multiple chromatin modifications can generate unanticipated stability of epigenetic States in Arabidopsis. Plant
Cell 22: 34–47.
132. Ha, M., D.W. Ng, W.H. Li & Z.J. Chen. 2011. Coordinated
histone modifications are associated with gene expression
variation within and between species. Genome Res. 21: 590–
598.
133. Ha, M., J. Lu, L. Tian, et al. 2009. Small RNAs serve as a
genetic buffer against genomic shock in Arabidopsis interspecific hybrids and allopolyploids. Proc. Natl. Acad. Sci. U.
S. A. 106: 17835–17840.
134. Kenan-Eichler, M., D. Leshkowitz, L. Tal, et al. 2011. Wheat
hybridization and polyploidization results in deregulation
of small RNAs. Genetics 188: 263–272.
135. Lackey, E., D.W. Ng & Z.J. Chen. 2010. RNAi-mediated
down-regulation of DCL1 and AGO1 induces developmental changes in resynthesized Arabidopsis allotetraploids.
New Phytol. 186: 207–215.
C 2014 New York Academy of Sciences.
Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 17
Plant polyploidy
Moghe & Shiu
136. Pikaard, C.S. & O. Mittelsten-Scheid. 2014. “Epigenetic
Regulation in Plants.” In Epigenetics. D. Allis, T. Jenuwein
& D. Reinberg, Eds. CSHL Press.
137. Coate, J.E., H. Bar & J.J. Doyle. 2014. Extensive translational
regulation of gene expression in an allopolyploid (Glycine
dolichocarpa). Plant Cell 26: 136–150.
138. Birchler, J.A. & R.A. Veitia. 2012. Gene balance hypothesis: connecting issues of dosage sensitivity across biological
disciplines. Proc. Natl. Acad. Sci. U. S. A. 109: 14746–14753.
139. De Smet, R. & Y. Van de Peer. 2012. Redundancy and
rewiring of genetic networks following genome-wide duplication events. Curr. Opin. Plant Biol. 15: 168–176.
140. Lou, P., J. Wu, F. Cheng, et al. 2012. Preferential retention
of circadian clock genes during diploidization following
whole genome triplication in Brassica rapa. Plant Cell 24:
2415–2426.
141. Bekaert, M., P.P. Edger, J.C. Pires & G.C. Conant. 2011.
Two-phase resolution of polyploidy in the Arabidopsis
metabolic network gives rise to relative and absolute dosage
constraints. Plant Cell 23: 1719–1728.
142. Veron, A.S., K. Kaufmann & E. Bornberg-Bauer. 2007. Evidence of interaction network evolution by whole-genome
duplications: a case study in MADS-box proteins. Mol. Biol.
Evol. 24: 670–678.
143. Hsu, C.Y., J.P. Adams, H. Kim, et al. 2011. FLOWERING
LOCUS T duplication coordinates reproductive and vegetative growth in perennial poplar. Proc. Natl. Acad. Sci. U.
S. A. 108: 10756–10761.
144. Sheehan, M.J., L.M. Kennedy, D.E. Costich & T.P. Brutnell.
2007. Subfunctionalization of PhyB1 and PhyB2 in the control of seedling and mature plant traits in maize. Plant J.
49: 338–353.
145. Wolfe, K.H. & D.C. Shields. 1997. Molecular evidence for
an ancient duplication of the entire yeast genome. Nature
387: 708–713.
146. Ihmels, J., S. Bergmann, M. Gerami-Nejad, et al. 2005.
Rewiring of the yeast transcriptional network through the
evolution of motif usage. Science 309: 938–940.
147. Conant, G.C. & K.H. Wolfe. 2006. Functional partitioning
of yeast co-expression networks after genome duplication.
PLoS Biol. 4: e109.
148. Levin, D. 2002. “Phenotypic consequences of chromosome
doubling.” In The Role of Chromosomal Change in Plant
Evolution. pp. 134–149. Oxford University Press.
149. Otto, S.P. & J. Whitton. 2000. Polyploid incidence and evolution. Annu. Rev. Genet. 34: 401–437.
150. Li, X., E. Yu, C. Fan, et al. 2012. Developmental, cytological
and transcriptional analysis of autotetraploid Arabidopsis.
Planta 236: 579–596.
151. Te Beest, M., J.J. Le Roux, D.M. Richardson, et al. 2012. The
more the better? The role of polyploidy in facilitating plant
invasions. Ann. Bot. 109: 19–45.
152. Robertson, K., E.E. Goldberg & B. Igić. 2011. Comparative
evidence for the correlated evolution of polyploidy and
self-compatibility in Solanaceae. Evolution 65: 139–155.
153. Barringer, B.C. 2007. Polyploidy and self-fertilization in
flowering plants. Am. J. Bot. 94: 1527–1533.
154. Mable, B.K. 2004. Polyploidy and self-compatibility: Is
there an association? New Phytol. 162: 803–811.
18
155. Warner, D.A. & G.E. Edwards. 1993. Effects of polyploidy
on photosynthesis. Photosynth. Res. 35: 135–147.
156. Li, W.L., G.P. Berlyn & P.M.S. Ashton. 1996. Polyploids and
their structural and physiological characteristics relative to
water deficit in Betula papyrifera (Betulaceae). Am. J. Bot.
U. S. A.
157. Coate, J.E., A.F. Powell, T.G. Owens & J.J. Doyle. 2013.
Transgressive physiological and transcriptomic responses
to light stress in allopolyploid Glycine dolichocarpa (Leguminosae). Heredity 110: 160–170.
158. Maherali, H., A.E. Walden & B.C. Husband. 2009. Genome
duplication and the evolution of physiological responses to
water stress. New Phytol. 184: 721–731.
159. Hao, G.-Y., M.E. Lucero, S.C. Sanderson, et al. 2013. Polyploidy enhances the occupation of heterogeneous environments through hydraulic related trade-offs in Atriplex
canescens (Chenopodiaceae). New Phytol. 197: 970–
978.
160. Levin, D.A. 1983. Polyploidy and novelty in flowering
plants. Am. Nat. 122: 1–25.
161. Oswald, B.P. & S.L. Nuismer. 2007. Neopolyploidy and
pathogen resistance. Proc. R. Soc. B 274: 2393–2397.
162. Schoen, D.J., J.J. Burdon & A.H. Brown. 1992. Resistance
of Glycine tomentella to soybean leaf rust Phakopsora
pachyrhizi in relation to ploidy level and geographic distribution. Theor. Appl. Genet. 83: 827–832.
163. Busey, P., R.M. Giblin-Davis & B.J. Center. 1993. Resistance
in Stenotaphrum to the sting nematode. Crop Sci. U. S. A.
164. Thompson, J.N., B.M. Cunningham, K.A. Segraves, et al.
1997. Plant polyploidy and insect/plant interactions. Am.
Nat. 150: 730–743.
165. Nuismer, S.L. & J.N. Thompson. 2001. Plant polyploidy
and non-uniform effects on insect herbivores. Proc. R. Soc.
B 268: 1937–1940.
166. Stebbins, C.L. 1950. Variation and Evolution in Plants. London: Oxford University Press.
167. Otto, S.P. 2007. The evolutionary consequences of polyploidy. Cell 131: 452–462.
168. Fawcett, J.A. & Y.V. de. Peer. 2010. Angiosperm polyploids
and their road to evolutionary success. Trends Evol. Biol. 2:
e3.
169. Brochmann, C., A.K. Brysting, I.G. Alsos, et al. 2004. Polyploidy in arctic plants. Biol. J. Linn. Soc. 82: 521–536.
170. Fawcett, J.A., S. Maere & Y. Van de Peer. 2009. Plants with
double genomes might have had a better chance to survive
the Cretaceous-Tertiary extinction event. Proc. Natl. Acad.
Sci. U. S. A. 106: 5737–5742.
171. Pandit, M.K., M.J.O. Pocock & W.E. Kunin. 2011. Ploidy influences rarity and invasiveness in plants. J. Ecol. 99: 1108–
1115.
172. Chao, D.-Y., B. Dilkes, H. Luo, et al. 2013. Polyploids exhibit higher potassium uptake and salinity tolerance in
Arabidopsis. Science 341: 658–659.
173. Bowden, W.M. 1940. Diploidy, polyploidy, and winter hardiness relationships in the flowering plants. Am. J. Bot. 27:
357–371.
174. Martin, S.L. & B.C. Husband. 2009. Influence of phylogeny and ploidy on species ranges of North American
angiosperms. J. Ecol. 97: 913–922.
C 2014 New York Academy of Sciences.
Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 Moghe & Shiu
175. Segraves, K.A. & J.N. Thompson. 1999. Plant polyploidy
and pollination: floral traits and insect visits to diploid and
tetraploid Heuchera grossulariifolia. Evolution 53: 1114–
1127.
176. Kennedy, B.F., H.A. Sabara, D. Haydon & B.C. Husband.
2006. Pollinator-mediated assortative mating in mixed
ploidy populations of Chamerion angustifolium (Onagraceae). Oecologia 150: 398–408.
177. Baldwin, S.J. & B.C. Husband. 2011. Genome duplication
and the evolution of conspecific pollen precedence. Proc.
R. Soc. B 278: 2011–2017.
178. Ainouche, M.L., P.M. Fortune, A. Salmon, et al. 2009. Hybridization, polyploidy and invasion: lessons from Spartina
(Poaceae). Biol. Invasions 11: 1159–1173.
179. Ramsey, J. 2011. Polyploidy and ecological adaptation in
wild yarrow. Proc. Natl. Acad. Sci. U. S. A. 108: 7096–7101.
180. Martin, S.L. & B.C. Husband. 2012. Whole genome duplication affects evolvability of flowering time in an autotetraploid plant. PLoS One 7: e44784.
181. Martin, S.L. & B.C. Husband. 2013. Adaptation of diploid
and tetraploid chamerion angustifolium to elevation but
not local environment. Evolution 67: 1780–1791.
182. Vamosi J.C. & T.A. Dickinson. 2006. Polyploidy and diversification: a phylogenetic investigation in Rosaceae. Int. J.
Plant Sci. 167: 349–358.
183. Paterson, A.H., J.E. Bowers & B.A. Chapman. 2004. Ancient
polyploidization predating divergence of the cereals, and its
consequences for comparative genomics. Proc. Natl. Acad.
Sci. U. S. A. 101: 9903–9908.
184. Barker, M.S., N.C. Kane, M. Matvienko, et al. 2008. Multiple paleopolyploidizations during the evolution of the
Compositae reveal parallel patterns of duplicate gene retention after millions of years. Mol. Biol. Evol. 25: 2445–2455.
Plant polyploidy
185. Cannon, S.B., D. Ilut, A.D. Farmer, et al. 2010. Polyploidy
did not predate the evolution of nodulation in all legumes.
PloS One 5: e11630.
186. Wood, T.E., N. Takebayashi, M.S. Barker, et al. 2009. The
frequency of polyploid speciation in vascular plants. Proc.
Natl. Acad. Sci. U. S. A. 106: 13875–13879.
187. Mayrose, I., S.H. Zhan, C.J. Rothfels, et al. 2011. Recently
formed polyploid plants diversify at lower rates. Science
333: 1257.
188. Sabara, H.A., P. Kron & B.C. Husband. 2013. Cytotype coexistence leads to triploid hybrid production in a diploidtetraploid contact zone of Chamerion angustifolium (Onagraceae). Am. J. Bot. 100: 962–970.
189. Marques, I., D. Draper, L. Riofrı́o & C. Naranjo. 2014. Multiple hybridization events, polyploidy and low postmating
isolation entangle the evolution of neotropical species of
Epidendrum (Orchidaceae). BMC Evol. Biol. 14: 20.
190. Sonnleitner, M., B. Weis, R. Flatscher, et al. 2013. Parental
ploidy strongly affects offspring fitness in heteroploid
crosses among three cytotypes of autopolyploid Jacobaea
carniolica (Asteraceae). PLoS One 8: e78959.
191. Petit, C., F. Bretagnolle & F. Felber. 1999. Evolutionary
consequences of diploid-polyploid hybrid zones in wild
species. Trends Ecol. Evol. 14: 306–311.
192. Ohta, T. 1992. The nearly neutral theory of molecular evolution. Annu. Rev. Ecol. Syst. 23: 263–286.
193. Umezawa, T., M. Fujita, Y. Fujita, et al. 2006. Engineering drought tolerance in plants: discovering and tailoring
genes to unlock the future. Curr. Opin. Biotechnol. 17: 113–
122.
194. Hu, H. & L. Xiong. 2013. Genetic engineering and breeding
of drought-resistant crops. Annu. Rev. Plant Biol. 65: 83–
108.
C 2014 New York Academy of Sciences.
Ann. N.Y. Acad. Sci. xxxx (2014) 1–19 19