Table of Contents
Cover
Title Page
Copyright
List of Contributors
Preface and Introduction
Acknowledgments
About the Editor
Part I: Somatic Genome Variation in Animals and Humans
Chapter 1: Polyploidy in Animal Development and Disease
1.1 Introduction
1.2 Mechanisms Inducing Somatic Polyploidy
1.3 The Core Cell Cycle Machinery
1.4 Genomic Organization of Polyploid Cells
1.5 Endoreplication: An Effective Tool for Post-Mitotic Growth and Tissue Regeneration
1.6 Initiation of Endoreplication in
Drosophila
1.7 Mechanisms of Endocycle Oscillations in
Drosophila
1.8 Gene Amplification in
Drosophila
Follicle Cells
1.9 Endocycle Entry in the Trophoblast Lineage
1.10 Mechanisms of Endocycle Oscillations in Trophoblast Giant Cells
1.11 Cardiomyocytes
1.12 Hepatocytes
1.13 Megakaryocytes
1.14 Concluding Remarks
Acknowledgments
References
Chapter 2: Large-Scale Programmed Genome Rearrangements in Vertebrates
2.1 Introduction
2.1 Hagfish
2.3 Sea Lamprey
2.4 Zebra Finch
2.5 Emerging Themes and Directions
References
Chapter 3: Chromosome Instability in Stem Cells
3.1 Introduction
3.2 Pluripotent Stem Cells
3.3 Somatic Stem Cells
3.3.1 Mesenchymal Stem Cells
3.4 Mechanisms of Chromosomal Instability
3.5 Mechanisms of Chromosomal Instability in Stem Cells
References
Part II: Somatic Genome Variation in Plants
Chapter 4: Mechanisms of Induced Inheritable Genome Variation in Flax
4.1 Introduction
4.2 Restructuring the Flax Genome
4.3 Specific Genomic Changes
4.4 What Happens When Plastic Plants Respond to Environmental Stresses?
4.5 When Do the Genomic Changes Occur and Are they Adaptive?
4.6 Is this Genomic Response of Flax Unique?
4.7 Concluding Remarks
Acknowledgments
References
Chapter 5: Environmentally Induced Genome Instability and its Inheritance
5.1 Introduction
5.2 Stress and its Effects on Genomes
5.3 Transgenerational Inheritance
5.4 Concluding Remarks
Acknowledgments
References
Chapter 6: The Mitochondrial Genome, Genomic Shifting, and Genomic Conflict
6.1 Introduction
6.2 Heteroplasmy and Sublimons
6.3 Cytoplasmic Male Sterility (CMS) in Plants
6.4 Mitochondrial Sublimons and CMS
6.5 Restorer Gene Evolution: Somatic Genetic Changes Drive Nuclear Gene Diversity?
6.6 Concluding Remarks
References
Chapter 7: Plastid Genome Stability and Repair
7.1 Introduction
7.2 Characteristics of the Plastid Genome
7.3 Replication of Plastid DNA
7.4 Transcription in the Plastid
7.5 The Influence of Replication and Transcription on Plastid Genome Stability
7.6 Plastid Genome Stability and DNA Repair
7.7 Outcomes of DNA Rearrangements
7.8 Concluding Remarks
References
Part III: Somatic Genome Variation in Microorganisms
Chapter 8: RNA-Mediated Somatic Genome Rearrangement in Ciliates
8.1 Introduction
8.2 Ciliates: Ubiquitous Eukaryotic Microorganisms with a Long Scientific History
8.3 Two's Company: Nuclear Dimorphism in Ciliates
8.4
Paramecium
: Non-Mendelian Inheritance Comes to Light
8.5
Tetrahymena
and the Origin of the scanRNA Model
8.6 Small RNAs in
Stylonychia
and
Oxytricha
8.7 Long Noncoding RNA Templates in Genome Rearrangement
8.8 Long Noncoding RNA: An Interface for Short Noncoding RNA
8.9 Short RNA-Mediated Heterochromatin Formation and DNA Elimination
8.10 Transposable Elements and the Origins of Genome Rearrangements
8.11 Transposons, Phase Variation, and Programmed Genome Engineering in Bacteria
8.12 Transposases, Noncoding RNA, and Chromatin Modifications in VDJ Recombination of Vertebrates
8.13 Concluding Remarks: Ubiquitous Genome Variation, Transposons, and Noncoding RNA
Acknowledgments
References
Chapter 9: Mitotic Genome Variations in Yeast and Other Fungi
9.1 Introduction
9.2 The Replication Process as a Possible Source of Genome Instability
9.4 Ploidy Maintenance and Chromosome Integrity Mechanisms
9.5 Concluding Remarks
References
Webliography
Part IV: General Genome Biology
Chapter 10: General Genome Biology 10: Genome Variation in Archaeans, Bacteria, and Asexually Reproducing Eukaryotes
10.1 Introduction
10.2 Chromosome Number in Prokaryote Species
10.3 Genome Size Variation in Archaeans and Bacteria
10.4 Archaeal and Bacterial Genome Size Distribution
10.5 Genomic GC Content in Archaeans, Bacteria, Fungi, Protists, Plants, and Animals
10.6 Correlation between GC Content and Genome or Chromosome Size
10.7 Genome Size and GC-Content Variation in Primarily Asexually Reproducing Fungi
10.8 Variation of Gene Direction
10.9 Concluding Remarks
Acknowledgments
References
Chapter 11: RNA Polyadenylation Site Regions: Highly Similar in Base Composition Pattern but Diverse in Sequence—A Combination Ensuring Similar Function but Avoiding Repetitive-Regions-Related Genomic Instability
11.1 General Introduction to Gene Number, Direction, and RNA Polyadenylation
11.2 Base Selection at the Poly(A) Tail Starting Position
11.3 Most Frequent Upstream Motifs in Microorganisms, Plants, and Animals
11.4 Motif Frequencies in the Whole Genome
11.5 The Top 20 Hexamer Motifs in the Poly(A) Site Region in Humans
11.6 Polyadenylation Signal Motif Distribution
11.7 Alternative Polyadenylation
11.8 Base Composition of 3′UTR in Plants and Animals
11.9 Base Composition Comparison between 3′UTR and Whole Genome
11.10 Base Composition of 3′COR in Plants and Animals
11.11 Base Composition Pattern of the Poly(A) Site Region in Protists
11.12 Base Composition Pattern of the Poly(A) Site Region in Plants
11.13 Base Composition Pattern of the Poly(A) Site Region in Animals
11.14 Comparison of Poly(A) Site Region Base Composition Patterns in Plants and Animals
11.15 Common U-A-U-A-U Base Abundance Pattern in the Poly(A) Site Region in Fungi, Plants, and Animals
11.16 Difference between the Most Frequent Motifs and Seqlogo-Showed Most Frequent Bases
11.17 RNA Structure of the Poly(A) Site Region
11.18 Low Conservation in the Overall Nucleotide Sequence of the Poly(A) Site Region
11.19 Poly(A) Site Region Stability and Somatic Genome Variation
11.20 Concluding Remarks
Acknowledgments
References
Chapter 12: Insulin Signaling Pathways in Humans and Plants
12.1 Introduction
12.2 Ranking of the Insulin Signaling Pathway and its Key Proteins
12.3 Diseases Caused by Somatic Mutations of the PI3K, PTEN, and AKT Proteins in the Insulin Signaling Pathway
12.4 Plant Insulin and Medical Use
12.5 Role of the Insulin Signaling Pathway in Regulating Plant Growth
12.6 Concluding Remarks
References
Chapter 13: Developmental Variation in the Nuclear Genome Primary Sequence
13.1 Introduction
13.2 Genetic Mutation, DNA Damage and Protection, and Gene Conversion in Somatic Cells
13.3 Programmed Large-Scale Variation in Primary DNA Sequences in Somatic Nuclear Genome
13.4 Generation of Antibody Genes in Animals through Somatic Genome Variation
13.5 Developmental Variation in Primary DNA Sequences in the Somatic Cells of Plants
13.6 Heritability and Stability of Developmentally Induced Variation in the Somatic Nuclear Genome in Plants
13.7 Concluding Remarks
References
Chapter 14: Ploidy Variation of the Nuclear, Chloroplast, and Mitochondrial Genomes in Somatic Cells
14.1 Introduction
14.2 Nuclear Genome in Somatic Cells
14.3 Plastid Genome Variation in Somatic Cells
14.4 Mitochondrial Genome in Somatic Cells
14.5 Organelle Genomes in Somatic Hybrids
14.6 Effects of Nuclear Genome Ploidy on Organelle Genomes
14.7 Concluding Remarks
Acknowledgments
References
Chapter 15: Molecular Mechanisms of Somatic Genome Variation
15.1 Introduction
15.2 Mutation of Genes Involved in the Cell Cycle, Cell Division, or Centromere Function
15.3 DNA Damage
15.4 Variation in Induction and Activity of Radical-Scavenging Enzymes
15.5 DNA Cytosine Deaminases
15.6 Variation in Protective Roles of Pigments against Oxidative Damage
15.7 RNA-Templated DNA Repair
15.8 Errors in DNA Repair
15.9 RNA-Mediated Somatic Genome Rearrangement
15.10 Repetitive DNA Instability
15.11 Extracellular DNA
15.12 DNATransposition
15.13 Somatic Crossover and Gene Conversion
15.14 Molecular Heterosis
15.15 Genome Damage Induced by Endoplasmic Reticulum Stress
15.16 Telomere Degeneration
15.17 Concluding Remarks
References
Chapter 16: Hypotheses for Interpreting Somatic Genome Variation
16.1 Introduction
16.2 Cell-Specific Accumulation of Somatic Genome Variation in Somatic Cells
16.3 Developmental Age and Genomic Network of Reproductive Cells
16.4 Genome Generation Cycle of Species
16.5 Somatic Genome Variation and Tissue-Specific Requirements during Growth or Development
16.6 Costs and Benefits of Somatic Genome Variation
16.7 Hypothesis on the Existence of a Primitive Stage in both Animals and Plants
16.8 Sources of Genetic Variation from in Vitro Culture Propagation
16.9 Hypothesis that Heterosis Is Created by Somatic Genome Variation
16.10 Genome Stability through Structural Similarity and Sequence Dissimilarity
16.11 Hypothesis Interpreting the Maternal Transmission of Organelles
16.12 Ability of Humans to Deal with Somatic Genome Variation and Diseases
16.13 Concluding Remarks
References
Chapter 17: Impacts of Somatic Genome Variation on Genetic Theories and Breeding Concepts, and the Distinction between Mendelian Genetic Variation, Somagenetic Variation, and Epigenetic Variation
17.1 Introduction
17.2 The Term ‘Somatic Genome’
17.3 Mendelian Genetic Variation, Epigenetic Variation, and Somagenetic Variation
17.4 What Is a Gene?
17.5 Breeding Criteria, Genome Cycle, Pure Lines, and Variety Stability
17.6 The Weismann Barrier Hypothesis and the Need for Revision
17.7 Implications for Species Evolution
17.8 Concluding Remarks
References
Chapter 18: Somatic Genome Variation: What it is and What it Means for Agriculture and Human Health
18.1 Introduction
18.2 Natural Attributes of Somatic Genome Variation
18.3 Implications of Somatic Genome Variation for Human and Animal Health
18.4 Implications of Somatic Genome Variation for Agriculture
18.5 Concluding Remarks
Acknowledgments
References
Index
End User License Agreement
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Guide
Cover
Table of Contents
Preface and Introduction
Part I: Somatic Genome Variation in Animals and Humans
Begin Reading
List of Illustrations
Chapter 1: Polyploidy in Animal Development and Disease
Figure 1.1 Mechanisms introducing somatic polyploidy. (A) Nuclear configurations in mitotic and polyploid cell types. C, C-value, multiples of the haploid DNA content arising from different mechanisms. (B) Overview of non-canonical cell cycle variants leading to somatic polyploidy. Hepatocytes (HPC) bypass cytokinesis, and as a consequence this cell cycle variant is called an acytokinetic mitosis. Megakaryocyte polyploidization occurs through endomitosis, which is another abortive mitosis lacking anaphase B, telophase, and cytokinesis. Mouse trophoblast giant cells (TGC) and
Drosophila
salivary glands (SG) both undergo endocycles, which are completely devoid of M phase. TGCs undergo full genome replications, while SGs exit S phase before completing late replication. (C) Schematic of an under-replicated region, which results from differential firing of replication origins and within the euchromatin usually encompasses 100–400 kb. (D) Genomic organization of amplified loci, which arise from reiterative origin firing, resulting in copy number gradients stretching approximately 100 kb.
Figure 1.2 Signaling pathways controlling cell cycle transitions in
Drosophila
ovarian follicle cells. (Top) A schematic depicts the development of the
Drosophila
egg chamber and delineates the points where follicle cells undergo cell cycle transitions. Each egg chamber consists of 16 germline cells, the diploid oocyte (red), and 15 polyploid nurse cells (orange), surrounded by a layer of somatic follicle cells (green). The follicle cells proliferate mitotically until stage 6, and then undergo three endocycles. At stage 10B, all genome-wide DNA replication ceases and amplification initiates at six specific loci. (Bottom left) Activation of the Notch signaling pathway in follicle cells is the major stimulus for the mitotic-to-endocycle transition. Notch signaling blocks mitosis in these cells by activating the transcriptional repressor Hnt. Hnt then represses the expression of Stg, as well as Cut, which triggers the degradation of the mitotic cyclins A and B by the APC/C–Fzr complex. In addition, Notch down-regulates the expression of Dap, preventing it from inhibiting S phase. CoREST is a transcriptional cofactor required for proper Notch signaling. (Bottom right) The endocycle-to-amplification transition requires the inactivation of Notch signaling to allow signaling through EcR. The crucial event downstream of both Notch silencing and EcR activation is increased expression of the transcriptional repressor Ttk69, which is required for the switch to amplification. miR-7 negatively regulates Ttk69 expression and can block amplification when overexpressed. Notch silencing also causes the down-regulation of Hnt expression, which allows Cut to be expressed. It is likely that additional signaling pathways are also involved in this cell cycle transition.
Figure 1.3 Endocycle oscillations. (A) Model of the autonomous oscillator that drives the salivary gland endocycles. The transcriptional activator E2F1 accumulates during the late G(ap) phase and stimulates CycE transcription, which triggers S phase in conjunction with its kinase partner Cdk2. Ongoing DNA replication activates the E3 ubiquitin ligase CRL4-Cdt2, which marks E2F1 for proteasomal degradation. The CRL1–Ago complex continuously targets CycE for degradation, and hence this negative feedback loop causes a drop in CycE levels, which is a prerequisite for origin licensing during the next G phase. CycE/Cdk2 also inhibits APC/C–Fzr activity, thereby allowing accumulation of the licensing inhibitor Geminin. (B) The regulatory network that controls the endocycle progression in trophoblast giant cells (TGC). S phase in TGCs relies on Cdk2, which associates with either CycE or CycA. During late G phase a peak of CycE/Cdk2 activity triggers S phase and concomitantly inhibits the ACP/C–Fzr complex, thereby promoting the accumulation of CycA and Geminin. CycA/Cdk2 phosphorylates CycE, which in turn will be recognized by E3 ligase CRL1-Fbw7 and thus marked for proteasomal degradation. The cessation of CycE/Cdk2 activity permits accumulation of CKI p57, which is no longer antagonized by the CRL1–Skp2 complex. The synergistic action of three factors, CRL1-Fbw7, CRL3, and p57, ensures low levels of CycE/Cdk2 activity during G phases, thereby allowing origin licensing to occur.
Figure 1.4 Abortive division cycles. (A) Schematic of normal mitotic cell division. During prophase, chromatin condenses into distinctive chromosomes and the mitotic spindle begins to assemble between duplicated centrosomes. Breakdown of the nuclear envelope marks the onset of prometaphase, thereby allowing spindle microtubules to connect with chromosomes. During metaphase, chromosomes align along the equator of the spindle and both of their kinetochores are attached to microtubules. At anaphase A, sister chromatids separate and are then pulled towards spindle poles by shortening of kinetochore microtubules. Anaphase B is characterized by lengthening of polar microtubules, thereby pushing spindle poles apart. In telophase (not shown) chromosomes decondense and the nuclear envelope starts to assemble. Finally, a contractile actin–myosin ring forms during cytokinesis and splits cytoplasm into two daughter cells. (B) During acytokinetic mitosis the mitotic program is executed normally, but cells fail to establish a contractile actin–myosin ring and cannot cleave, which leads to formation of binucleate cells with two centrosomes. (C) Megakaryocytes undergoing endomitosis normally progress through prophase, prometaphase, metaphase, and anaphase A, but fail to execute anaphase B, telophase, or cytokinesis. As a consequence, the reassembling nuclear envelope encloses the sister chromatids in a single nucleus, thereby giving rise to a characteristic lobular structure. Higher levels of ploidy are accomplished by consecutive rounds of endomitosis, which involve centrosome duplication and formation of multipolar spindles. (
See plate section for color representation of this figure.
)
Chapter 2: Large-Scale Programmed Genome Rearrangements in Vertebrates
Figure 2.1 Distribution of germline-specific DNA and patterns of DNA elimination across the vertebrate phylogeny. Germline-specific sequences have been observed in lamprey, hagfish, and zebra finch, and studies have begun to reveal the causes and consequences of programmed genome rearrangement in these groups. It is not known if hagfish and lamprey retain programmed genome rearrangement as an aspect of their biology inherited from their common ancestor or if similar genome biologies evolved more than once in the two lineages. The germline-specific chromosome of zebra finch appears to have arisen relatively recently and likely represents an independent evolutionary acquisition of germline-specific DNA. Most vertebrate lineages have not been rigorously tested for the presence or absence of germline-specific DNA. MYA, Million years ago.
Chapter 3: Chromosome Instability in Stem Cells
Figure 3.1 Mitotic spindle organization: (A) normal bipolar spindle with two centrosomes; (B) multipolar spindle with three centrosomes; (C) bipolar spindle with clustered supernumerary centrosomes. In SCs only multipolar spindles with three or more centrosomes were described. Light gray bars represent mitotic chromosomes.
Chapter 4: Mechanisms of Induced Inheritable Genome Variation in Flax
Figure 4.1 Induction of heritable changes in flax. The growth of the original line Stormont Cirrus (Pl) under different environmental conditions can result in new stable phenotypes with associated genomic variation. Shown is the appearance of the small genotroph following growth for a single generation under inducing conditions, and individuals after 40 additional generations of growth under standard non-inducing conditions where the original phenotypic and genotypic differences have persisted.
Figure 4.2 Alignment of next-generation sequencing reads (approximately 100× sequencing) from Pl, and the two small genotrophs S and C3, to the Bethune genome. Both S and C3 have a deletion in a region where Pl has the same sequence as Bethune. The deletion has been confirmed by PCR across the region, where the lane MI is Bioline molecular marker I.
Figure 4.3 Alignment of next-generation sequencing reads (approximately 100× sequencing) from Pl, and the two small genotrophs S and C3, to the Bethune genome. Pl differs from the genotrophs with a region not present in Pl but present in the genotrophs, and that region in the genotrophs is identical to the sequence in Bethune. The deletion has been confirmed by PCR across the region, where the lane MI is Bioline molecular marker I.
Figure 4.4 Alignment of next-generation sequencing reads (approximately 100× sequencing) from Pl, and the two small genotrophs S and C3, to the Bethune genome. There are copy number differences between Pl and the genotrophs. The number of reads from the Pl sample is five to six times greater than that from either S or C3. S and C3 appear slightly different from each other, which has previously been observed in the genotrophs where the 5S and large rRNA genes vary but were not exactly identical in all lines.
Figure 4.5 Changes in LIS-1 and 5SrDNA following growth under various nutrient regimes. PCR amplifications from DNA extracted from the upper leaves of Stormont Cirrus plants grown under three different nutrient regimes, with primers (a) spanning the left junction of LIS1, (b) the uninserted site, (c) the right junction of LIS1, and (d) primers specific for the spacer region of 5SrDNA represented by the clone pBG13 (Goldsbrough and Cullis 1981). The two bands in (d) represent the two spacer length variants found in pBG13 and differ by 21 base pairs. Lanes 1 and 2 show two plants grown with an imbalance of nitrogen; lanes 3 and 4, two plants grown in soil without any added nutrients; lanes 5 and 6, two plants grown with balanced nutrients. M, Molecular weight marker VI (Roche). All the other plants grown under each of the set of nutrient conditions had the same LIS-1 status as those shown in the Figure However, there was no consistent pattern of relative amounts of the two spacer-length variants for the 5SrDNA among these same plants. This complete reproducible conversion of all the plants to homozygous LIS-1 containing under two of the nutrient regimes and never under the third regime is consistent with LIS-1 being selected for under some nutrient regimes and either neutral or deleterious under others.
Chapter 5: Environmentally Induced Genome Instability and its Inheritance
Figure 5.1 A hypothetical scheme connecting environmental stress and epigenetic/genetic changes that promote soft and hard (Mendelian) inheritance.
Chapter 6: The Mitochondrial Genome, Genomic Shifting, and Genomic Conflict
Figure 6.1 Genetic events implicated in the occurrence of gynodioecy in plant populations. In a population of hermaphroditic plants (open symbols), the appearance of a new mitochondrial CMS mutation (1, filled, female symbols) can spread due to its maternal mode of inheritance, resulting in a rise in the proportion of female, male sterile plants (2). This eventually places a limitation on pollen production that creates the selective force to drive a response by the nuclear genome, the evolution of a new fertility restorer gene (3). This suppresses the male sterile trait, and leads to the appearance of hermaphroditic plants with the mitochondrial male sterility gene (filled hermaphroditic symbols). In the case shown here, the new CMS–restorer combination eventually spreads to fixation (4), masking the presence of the CMS mitochondrial genome.
Chapter 7: Plastid Genome Stability and Repair
Figure 7.1 The D-loop plastid DNA replication model. In (a) to (f), black lines represent the two parental strands. Red and blue lines represent the two daughter leading strands and lagging strands, respectively. In (a) to (g), arrows represent direction of DNA polymerization (3′ end). (
See plate section for color representation of this figure.
)
Figure 7.2 Rolling circle plastid DNA replication. Plain lines represent parental strands. Dotted and gray lines represent leading and lagging strands, respectively. Arrows represent direction of DNA polymerization (3′ end).
Figure 7.3 ROS generation and scavenging. Energy or electron transfer to molecular oxygen generates, respectively, two types of ROS: singlet oxygen and superoxide anions. Superoxide dismutases and ascorbate peroxidases constitute the main scavenging process to eliminate superoxide and avoid production of highly reactive hydroxyl radical.
1
O
2
, Singlet oxygen;
3
O
2
, dioxygen; e
–
, electron; O
2
–
, superoxide anion; SODs, superoxide dismutases; H
2
O
2
, hydrogen peroxide; APXs, ascorbate peroxidases; •OH, hydroxyl radical;
–
OH, hydroxide anion.
Figure 7.4 Oxidation of guanine and thymine mainly leads to generation of 8-oxoguanine and thymine glycol, respectively.
Figure 7.5 UV-B-induced linkage of adjacent pyrimidines mainly leads to formation of cyclobutane pyrimidine dimers (CPDs) and 6-4 photoproducts.
Figure 7.6 Repair of a DSB by homologous recombination. Dotted lines represent newly synthesized DNA and purple lines represent stretches of homologous sequence. SSA, Single-strand annealing; SDSA, synthesis-dependent strand annealing; DSBR, double-strand break repair.
Source
: Adapted from Hastings et al. 2009b. (
See plate section for color representation of this figure.
)
Chapter 8: RNA-Mediated Somatic Genome Rearrangement in Ciliates
Figure 8.1 Simplified
Oxytricha
lifecycles. A: Reproductive asexual cycle. Cells divide by mitosis of the diploid germline micronucleus (MIC, indicated by a circle) and amitosis of the polyploid somatic macronucleus (MAC, indicated by an oval). B: In the laboratory, starvation induces conjugation between compatible mating types and initiation of a non-reproductive sexual cycle. C: Meiosis of the MIC produces haploid gametic nuclei. D: Exchange of haploid micronuclei occurs. E: Fertilization produces genetically distinct, diploid zygotic nuclei. F: Zygotic nuclei divide mitotically and one differentiates into a new MAC, while one remains as a new MIC. G: Old MACs shrink as their genomic content is degraded, and these late-stage cells are known as donut cells due to the very large, relatively transparent, developing zygotic MAC. H: At completion of the genome rearrangement process, the newly formed reproductive cell proliferates asexually (A) or can conjugate again (after a latency period), depending on nutrient availability.
Figure 8.2 Summary of noncoding RNA (ncRNA) in ciliate genome rearrangements. Panels A–C represent nuclei simultaneously present in one
Paramecium
or
Tetrahymena
cell, while panels D–F represent nuclei present in one
Oxytricha
cell. A–C: ScanRNA model in
Paramecium
and
Tetrahymena
. ScanRNA biogenesis initiates in the micronucleus (MIC) (A), where all sequences are transcribed into long noncoding ncRNAs that are subsequently processed by Dicer-like proteins into scanRNAs. B: Small RNAs are scanned against the parental macronuclear (MAC) genome via RNA–RNA interaction with long ncRNAs. Those that match are eliminated (indicated by scissors symbol) and only non-matching scan RNAs are retained for use in the developing zygotic MAC (C). C: Elimination of IES sequence is accomplished by hybridization of scanRNAs to nascent ncRNAs which target chromatin modifying enzymes to eliminated sequences. D–F: piRNA and template model developed in
Oxytricha.
D: The MIC genomic sequence remains quiescent. E: Transcription of all MAC chromosomes produces templates covering all genomic sequence (labeled ‘templates’). PiRNAs are also produced, either from these sequences (as shown) or from independent precursor molecules (not shown). Both classes of noncoding RNA are imported into the zygotic MAC (F), where they play different functional roles. F: PiRNAs pair with nascent ncRNAs (produced by transcription of precursor DNA molecules) and direct retention of genomic sequences. Template RNA molecules guide segment reordering and ligation of cleaved DNA. In all panels, orange represents RNA, while DNA is indicated as a black double helix. Blue indicates the sequence to be eliminated in the new MAC, and can be encoded in DNA or RNA depending on species and developmental stage.
Figure 8.3 Experimental detection of noncoding RNA (ncRNA) from eliminated micronuclear (MIC) regions during sexual cycle in
Oxytricha
(corresponding to the nascent ncRNAs shown in Figure 8.2F)
.
An agarose northern membrane was probed with a MIC-limited 170-bp satellite repeat revealing noncoding RNAs ranging from several hundred basepairs to over 10,000 bp. The abundance of these ncRNAs peaks at 18 h and 24 h post mixing of complementary mating types JRB310 and JRB510. The membrane was stripped and reprobed with
Actin1
as a loading control, and ethidium-stained ribosomal RNAs (28S and 18S) from the gel prior to transfer are shown as additional load controls due to expression fluctuation in
Actin1
during the sexual cycle.
Figure 8.4 Overview of autonomous and non-autonomous transposable elements in ciliates and other eukaryotes. Black boxes indicate telomeres, which are added de novo to ends of macronuclear (MAC) chromosomes, after micronuclear (MIC) chromosome fragmentation. Light-green regions are either mobile elements or thought to have mobile element origins (e.g., IESs). Arrowheads on both ends of most green sequences represent inverted terminal repeats. Blue arrows indicate ORFs encoding transposase enzyme. Dark-green triangles indicate direct repeats. Curved arrows indicate excision or mobilization activity of transposase enzymes. A: IES excision in
Euplotes crassus.
MIC transposase-encoding Tc1/mariner elements (Tec elements) are thought to control excision of IESs (though experimental proof is lacking so the arrows are dashed). Tec elements and IESs are flanked by TA motifs and the same inverted terminal repeats. B: IES excision in
Oxytricha trifallax
. MIC-encoded Tc1/mariner elements (telomere-bearing elements, or TBEs) produce transposase required for excision of IESs. TBE elements are flanked by ANT motifs and strong (72-bp) inverted terminal repeats, while IESs have extremely weak inverted terminal repeat tendencies only detectable in aggregate (Chen et al. 2014). IESs are generally flanked by direct repeats of 2–20 bp known as
pointers
; the 2-bp-long instances are biased toward TA (Chen et al. 2014). C: IES excision in
Paramecium tetaurelia.
A MAC gene (
PiggyMac
(
PGM
)) produces a PiggyBac family transposase required for IES excision. IESs are always flanked by TA and have inverted terminal repeats similar to the Euplotes Tec and IES motif. D: IES excision in
Tetrahymena thermophila
. As in
Paramecium
, a MAC gene (
Tetrahymena PiggyBac 2
(
TBP2
)) produces PiggyBac family transposase that is required for IES excision. Most IESs are long (>500 bp), lack inverted terminal repeats and direct repeats, and imprecisely excised; however, a small number of IESs (not shown in figure) are short (< 500 bp), flanked by TTAA, and precisely removed (Fass et al. 2011). E: Many eukaryotes have both autonomous and non-autonomous transposable elements analogous to ciliate transposon/IES systems. Autonomous elements are self-mobilizing and encode functional transposase enzyme, while non-autonomous versions (miniature inverted-repeat transposable elements (MITEs)) lack the coding sequence for functional transposase and therefore rely upon the enzyme produced from autonomous elements to mediate their transposition.
Source
: Chen et al., 2014; Fass et al. 2011; Smit and Riggs 1996; Casacuberta and Santiago 2003; Wessler 2006; Yang et al. 2009. Reproduced with permission of Springer. (
See plate section for color representation of this figure.
)
Figure 8.5 An example of developmentally programmed genome rearrangement in a prokaryote. The cyanobacteria
Anabaena
contains two operons with genes necessary for nitrogen fixation (shown in B). Both operons are interrupted by large (11- and 55-kb) insertions that are developmentally excised upon differentiation into heterocysts and resemble ciliate IESs, since they interrupt protein-coding regions. A:
Anabaena
vegetative cell DNA contains the insertion elements. Open triangles represent direct repeats of 5′-GCCTCATTAGG-3′ occurring at breakpoint for the 11-kb programmed deletion; filled triangles represent direct repeats of 5′-TATTC-3′ which demarcate boundaries of the 55-kb element. As with
Oxytricha
(pointers) and direct repeats in other ciliates, one copy of the repeat is retained in the rearranged DNA molecule.
XisA
and
xisF
denote recombinases responsible for removal of 11-kb and 55-kb segments, respectively, and which are not present in product (heterocyst) DNA. B: Structure of heterocyst DNA containing two intact operons (shown as arrows underneath the DNA sequence). Both sequence insertions interrupt coding regions and each insertion inhibits expression of one operon in vegetative cells.
Source
: Redrawn from Haselkorn 1992.
Chapter 9: Mitotic Genome Variations in Yeast and Other Fungi
Figure 9.1 DNA perturbations, triggered by various events, induce a multitude of molecular mechanisms that lead to damage repair, mutagenesis, or cell death.
Figure 9.2 Controlling of DNA repair pathways by various covalent modifications of PCNA.
Figure 9.3 Causes and consequences of cellular DNA content disturbances.
Chapter 10: General Genome Biology 10: Genome Variation in Archaeans, Bacteria, and Asexually Reproducing Eukaryotes
Figure 10.1 Average genome size of archaeans, gram-positive bacteria, and gram-negative bacteria. The number of genomes is 61,184, and 235 for archaeans, gram-positive bacteria, and gram-negative bacteria, respectively. The mean genome size order is 1.77 Mb < 2.53 Mb < 2.67 Mb for archaeans, gram-positive bacteria, and gram-negative bacteria, respectively. Note that both bacteria and archaeans showed considerable variation in genome size.
Figure 10.2 Normal probability plots of genome/chromosome size (bp, the Y-axis). Note that the large genomes/chromosomes (on the right side of each plot) do not fit a normal distribution and that archaeal genomes form two large groups.
Figure 10.3 The genomic base compositions and C+G contents of different kingdoms and large groups. The information is provided by phylum/class for archaeans, gram-positive bacteria, gram-negative bacteria, fungi, and protists and by species for dicot plants, monocot plants, non-mammalian animals, non-primate mammalian animals, and primate animals. Note that (1) C+G content is much more variable in prokaryotes than in higher organisms, (2) among plants and animals, C+G content is highest in monocot plants and lowest in dicot plants, and (3) mammals have very similar C+G contents between species.
Figure 10.4 Ratio (fold) of highest genomic GC content and lowest genomic GC content in prokaryotes and of the highest chromosomal GC content and lowest chromosome GC content in eukaryotes. GramP, Gram-positive bacteria; GramN, gram-negative bacteria. Average GC-content difference between prokaryotes and eukaryotes is highly significant (t-test,
P
< 0.01). Note that the genomic (or chromosomal for eukaryotes) GC-content ratio between the highest GC content and the lowest GC content is significantly higher in prokaryotes.
Figure 10.5 Distribution of values of chromosome size and C+G content in bacteria, archaeans, fungi, and protists. The chromosome number for bacteria, archaea, fungi, and protists is 430, 61, 139, and 21, respectively. Note that C+G content and chromosome or genome size is positively correlated in bacteria, but there is no such correlation or linear relationship in other kingdoms.
Figure 10.6 (A) Genome size and (B) GC contents of primarily asexual fungi and the fungi that are known to have a sexual reproductive stage. NCBI BioProjects ID for the primarily asexually producing fungi: PRJNA192877, PRJNA242986, PRJNA242987, PRJNA245139, PRJNA245140, PRJNA79339, RJNA221524, PRJNA67299, PRJNA174039, PRJNA73163, PRJNA10623, PRJNA78153, PRJNA39551, PRJNA18881, and PRJNA79337, for the species listed in this figure, from left to right, respectively. Species name (in order from left to right):
Hirsutella thompsonii
,
Hypocrella siamensis
,
Isaria farinose
,
Metarhizium acridum
,
Metarhizium anisopliae
,
Myceliophthora thermophila
,
Neotyphodium aotearoae
,
Neotyphodium gansuense
(1),
Neotyphodium gansuense
var.
inebrians
(2),
Tolypocladium inflatum
,
Ashbya gossypii
,
Eremothecium cymbalariae
,
Saccharomycetaceae
sp. ‘
Ashbya aceri
’,
Scheffersomyces stipites
, and
Thielavia terrestris.
Note that genome size is larger on average in the primarily asexually reproducing fungi than in the partly sexually reproducing fungi.
Chapter 11: RNA Polyadenylation Site Regions: Highly Similar in Base Composition Pattern but Diverse in Sequence—A Combination Ensuring Similar Function but Avoiding Repetitive-Regions-Related Genomic Instability
Figure 11.1 Poly(A) site region, presented using a UC poly(A) site as an example. In this illustration, the poly(A) tail attachment position (position −1) is a thymine. The poly(A) tail starting position (position 1) is a cytosine (C). In this chapter, we focus on the region from −100 to 100, which is a 201-base region. The poly(A) tail starting position is often loosely called ‘the poly(A) site’ in the literature. Note that although this illustration uses a C for the poly(A) tail starting position, in most species an adenine (A) occupies this position.
Figure 11.2 C/G ratios at the poly(A) tail attachment position of non-A-type poly(A) transcripts. Species were sorted from smallest to largest (from left to right) by C/G ratio at the poly(A) tail attachment position, giving the following ranking: dog, rabbit, rat, zebrafish, mouse, cattle, zebra finch, orangutan, chicken, human, pig, fruit fly. The three dicot plants were ranked as follows:
M. truncatula
,
A. thaliana
, poplar. The order for the three monocot plants was rice, maize, and sorghum. Note that the C/G ratio at the poly(A) tail attachment position clearly separates the three groups: animals < dicots < monocots (Li and Du 2013). (
See plate section for color representation of this figure.
)
Figure 11.3 Most frequent 20 hexamer motifs in the poly(A) site region of 30,499 unique poly(A) sites mapped to the human genome. Order of peak point labeling: motif sequence, position, and frequency at a single specific position in the mapped poly(A) site regions (pre-mRNA). The poly(A) sites are unique because each group of identical poly(A) sites was counted as one, which means that each of the 30,499 poly(A) sites was unique. Note that the most frequent motifs are mainly in three places: the −21 position region (upstream A-rich element; mainly AAUAAA), the poly(A) site, and the downstream U-rich region with peak location approximately at +19. Note also that the five most abundant motifs in the 201 bases (including 3′UTR, the poly(A) site, and 3′COR) in terms of values of highest peaks were AAUAAA, CAAAAA, UAAAAA, UUAAAA, and AAAUAA (Li and Du 2014b). (
See plate section for color representation of this figure.
)
Figure 11.4 Motif location around poly(A) sites. Frequency (%) is the percentage of mRNAs that have the motif's first nucleotide starting at the poly(A) site position. The UUACUU motif in
Trypanosoma cruzi
started at position −2 (with the nucleotide A directly at the poly(A) site). The UGUAAC motif in
Chlamydomonas reinhardtii
was at position −18 (i.e., the poly(A) site and the motif's last base were separated by 12 bases). The UGUUUG motif in
Blastocystis hominis
was located at downstream position 5 (i.e., the poly(A) tail attachment position and the motif's first base were separated by four bases). The UAUUUU motif was not very concentrated at any given location. The AAUAAA motif had its peak value at position −21 in humans and most animals, at position −22 in
Arabidopsis
(data not shown), and at position −24 in maize (
Zea mays
) (but the peak was small) (Li and Du 2014b).
Figure 11.5 RNA base compositions of the 3′UTR region, represented by 100 bases upstream of the poly(A) site. (a) Base compositions of 3′UTR and whole genome of each species. (b) 3′UTR/genome ratios of base composition. Note that the U-content difference between dicotyledonous and monocotyledonous plants is smaller in the 3′UTR region than in the whole genome, likely due to a higher overrepresentation of T content in terms of UTR/genome ratio in monocotyledonous plants (Li 2014). (
See plate section for color representation of this figure.
)
Figure 11.6 U-content overrepresentation in terms of 3′COR/genome ratio in comparison between dicotyledonous and monocotyledonous plants. 3′COR, The three-prime cleaved-off region downstream of the poly(A) site. Mapping was performed using mRNA sequences from the NCBI database. Note that monocotyledonous plants (the upper line) showed a greater 3′COR/genome ratio for U frequency at every position of the region than did dicotyledonous plants (the lower line) (Li 2014).
Figure 11.7 Average base abundance at each position of the 45 nucleotide poly(A) site regions in protists. Seqlogo position from left to right: 5′ to 3′ ends of the sequences. In these seqlogo graphs, position 31 is the poly(A) tail starting position (usually called ‘the poly(A) site’). Note that (1) the pattern of base abundance is very different in each of the protist species; (2) the poly(A) site is mainly an A in
Chlamydomonas reinhardtii
,
Phytophthora infestans
, and
Trypanosoma cruzi
, but is not an A in
Blastocystis hominis
; (3) the poly(A) tail attachment position is mainly a G for
B. hominis
and
C
.
reinhardtii
but a T in
T
.
cruzi
and a C or a T at approximately similar frequency for
P
.
infestans
; and (4) the signature region could be upstream of (
C
.
reinhardtii
), at (
T
.
cruzi
), or downstream of (
B
.
hominis
) the poly(A) site (Li and Du 2014b).
Figure 11.8 Base composition pattern around poly(A) sites in plants. Seqlogo graphs were produced based on average base composition among mapped poly(A) sites. Position 31 in these graphs is the poly(A) tail starting position, and position 30 is the poly(A) tail attachment position. Note that base abundance regions showed a U-A-U-A-U pattern (from left to right), with the second A corresponding to the poly(A) site (Li and Du 2014b).
Figure 11.9 Base composition pattern around poly(A) sites in various animal species:
Apis mellifera
(honey bee);
Drosophila melanogaster
(common fruit fly);
Taeniopygia guttata
(zebra finch);
Mus musculus
(mouse);
Homo sapiens
(human). Seqlogo graphs were produced based on average base composition among mapped poly(A) sites in each species. Position 31 in these graphs is the poly(A) tail starting position. Note that (1) the poly(A) site showed a strong A predominance, and (2) the predominant base at the poly(A) tail attachment position was U or C, depending on species (Li and Du 2014b).
Figure 11.10 Average base abundance at each position of the 201-nucleotide poly(A) site region, showing the U-A-U-SiteA-U (U-A-U-A-U for short) base abundance pattern in plants and animals. Species for plants:
Arabidopsis thaliana
,
Medicago truncatula
,
Oryza sativa
,
Populus trichocarpa
,
Solanum lycopersicum
,
Sorghum bicolor
,
Zea mays
. Species for animals:
Bos taurus
,
Danio rerio
,
Drosophila melanogaster
,
Gallus gallu
s,
Homo sapiens
,
Mus musculu
s,
Pongo abelii
,
Rattus norvegicus
,
Taeniopygia guttata
. Position 1 is the poly(A) tail starting position (usually called ‘the poly(A) site’). Position −1 is the poly(A) tail attachment position. Note that base composition analysis showed several clear differences between plants and animals. For example, the upstream A peak was lower but the U peak was higher in plants in comparison with animals. The shape of the downstream U peak was flat in plants but more pointed in animals. A small A peak at position −3 in plants and at −5 in animals exists, in addition to the U-A-U-SiteA-U (or U-A-U-A-U) base abundance pattern. Clearly, the U-A-U-A-U base abundance pattern is more conserved than any known motifs (Li and Du 2014b). (
See plate section for color representation of this figure.
)
Figure 11.11 Patterns of average base composition around poly(A) sites in fungi, plants, and animals. Seqlogo graphs were produced based on average base composition among mapped poly(A) sites. Position 31 in these graphs is the poly(A) tail starting position (corresponding to position 1 in Figure 11.1) and position 30 is the poly(A) tail attachment position (corresponding to position −1 in Figure 11.10). Positions 13 and 22 in these seqlogo graphs correspond to positions −18 and −9 in Figure 11.10. Note that base composition pattern was highly conserved among the fungal, plant, and animal kingdoms (Li and Du 2014b).
Figure 11.12 Predicted RNA folding structure of the poly(A) site region. (A) AB061250.1 (potato). (B) EU961162.1 (maize). 3′COR sequences were from their corresponding genes. Structures were determined by the MFOLD software program (http://unafold.rna.albany.edu/) using 201-bp poly(A) site region sequences that included 100 bp of the 3′UTR, poly(A) site, and 3′COR. Star: poly(A) site (phosphodiester bond) (note that the site is in a loop). Arrow: AAUAAA motif.
Chapter 12: Insulin Signaling Pathways in Humans and Plants
Figure 12.1 Brief presentation of the insulin signaling pathway, its connecting or overlapping signaling pathways, and the cellular processes directly affected by the pathways. AGGF1, angiogenic factor with G patch and FHA domains 1; AKT, protein kinase B; CBL, casitas B-lineage lymphoma, an E3 ubiquitin-protein ligase; FAS, fatty acid synthase; GYS, glycogen synthase; GLUT4, glucose transporter type 4; MAPK, mitogen-activated protein kinase; mTOR, mammalian target of rapamycin (serine/threonine kinase); PDK, phosphoinositide-dependent protein kinase; αPKC, protein kinase C, alpha; PI3K, phosphoinositide 3 kinase; PTEN, phosphatase and tensin homolog; PIP2, phosphatidylinositol 4,5-bisphosphate or PtdIns(4,5)P2; PIP3, phosphatidylinositol 3,4,5 trisphosphate; RhebGTP/RhebGDP, GTP-bound state or GDP-bound state, respectively, of Ras homolog enriched in brain, a GTPase; SREBP, sterol regulatory element-binding protein, a transcription factor; TSC, tuberous sclerosis. Note that this pathway is involved in carbohydrate metabolism, fatty acid biosynthesis, protein synthesis, and cellular proliferation.
Chapter 13: Developmental Variation in the Nuclear Genome Primary Sequence
Figure 13.1 A somatic crossover spot showing a dark-green region and an albino portion side by side on a leaf of a semi-dominant yellow mutant in
Nicotiana sylvestris
. Arrows indicate the green (dark) and yellow (light) twin spots.
Source
: From Li 1987.
Figure 13.2 Relative amount of DNA per root-cell nucleus among wild-type
Nicotiana sylvestris
and its doubled-haploid plants regenerated from microspore culture. Amount of DNA per nucleus was estimated by Feulgen cytophotometry of root-tip squashes.
Source
: Adapted from Li 1983.
Chapter 14: Ploidy Variation of the Nuclear, Chloroplast, and Mitochondrial Genomes in Somatic Cells
Figure 14.1 Karyotype showing maize cell ploidy levels under a microscope. (a) A diploid root cell (2n = 2x = 20); (b) a diploid leaf cell (2n = 2x = 20); and (c) a polyploid leaf cell (2n = 4x = approx. 40). The arrow in (a) points to two B-chromosome-like structures.
Source
: From Ma and Li 2015. (
See plate section for color representation of this figure.
)
Figure 14.2 Approximate copy number of the plastid genome per cell, estimated using real-time qPCR in seedlings of two diploid maize (
Zea mays
L.) cultivars, assuming that the leaves and roots were chiefly diploid. A single-copy, unique sequence of the nuclear genome was used as the copy number control. If values on the y-axis are divided by two (the assumed ploidy level), the values become the ratio of plastid DNA to nuclear DNA.
Source
: Adapted from Ma and Li 2015.
Chapter 17: Impacts of Somatic Genome Variation on Genetic Theories and Breeding Concepts, and the Distinction between Mendelian Genetic Variation, Somagenetic Variation, and Epigenetic Variation
Figure 17.1 Three categories of genome variation: Mendelian genetic variation (approximately equivalent to meiotic genetic variation), somagenetic variation, and epigenetic variation. Mendelian genetic variation is mainly meiotic genetic variation that follows the laws governing chromosome segregation and transmission between sexual generations. Somagenetic variation includes somatic genome variation as well as non-Mendelian genetic variations of the germline genome. Epigenetic variation includes DNA methylation and changes in gene expression; ‘epi’ means not the genome primary sequence itself. The three categories of variation interact and are frequently involved in the same biological processes.