2.1. Comparative Genomicsshodhganga.inflibnet.ac.in/bitstream/10603/12273/5/08_chapter 2.pdf · 2.1. Comparative Genomics In the post-genomic era, comparative analyses between genomes

Chapter 2 Review of Literature

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2.1. Comparative Genomics

In the post-genomic era, comparative analyses between genomes have proved

invaluable. It has emerged as a powerful tool to decipher gene and genome evolution

and improve genome annotation. The availability of fully annotated genome

sequence, tools and resources for functional genomics of Arabidopsis thaliana may be

helpful for comparative genome studies in crop plants. Multiple species comparison

has revealed novel insight into genome evolution (Bennetzen 2005; Donoghue et al.

2011), genome duplication (Ilic et al. 2003; Cheung et al. 2003; Parkin et al. 2005)

origin of new gene (Yang et al. 2008a), and can also identify unknown or poorly

characterized genome components, such as novel transposable and functional

elements (Jiang et al. 2004; Lai et al. 2005; Jiang et al. 2011). The complete genome

sequence of A. thaliana (Arabidopsis Genome Initiative 2000) provides a concrete

support for analysis of outcome of the diploidization process not only at the sequence

level directly within the genome of Arabidopsis (Blanc et al. 2000; Paterson et al.

2000; Paterson et al. 2001), but also in relation to sequences from distantly related

species, including Solanum lycopersicum (Ku et al. 2000), Oryza sativa (Yu et al.

2005a; Matsumoto et al. 2005), Sorghum bicolor (Paterson et al. 2009) and Maize

(Schnable et al. 2009). The National Science Foundation (NSF) launched the

Arabidopsis 2010 project with an aim to determining the function of ca. 25,000 genes

of Arabidopsis by 2010 (Somerville and Dangl 2000). Phylogenetic studies placed the

genus Brachypodium closer to rice (Catalan and Olmstead 2000). Therefore,

Brachypodium species were suggested to bridge the ‘genomic gap’ between rice and

the Triticeae family which contains some of the world’s most important crops

including wheat and barley (Draper et al. 2001; Foote et al. 2004). The angiosperm

family Brassicaceae contains both the research model A. thaliana and the agricultural

genus Brassica. Comparative genomics in the Brassicaceae has largely focused on

direct comparison between Arabidopsis and the species of interest. The discrimination

and appreciation of whole genome duplication (polyploidy) within lineages is crucial

for comparative studies within the Brassicaceae (Marhold and Lihova 2006).

Comparative genomics of A. thaliana and Brassica species is of importance for

understanding the evolution of their genome and to isolate and characterize the loci of


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interest. Early comparative studies conducted at the level of genetic linkage maps

revealed extensive duplication within Brassica genome (Lagercrantz and Lydiate

1996) and tracts of collinearity disrupted by multiple rearrangements between the

genomes of B. nigra and A. thaliana (Lagercrantz 1998). Subsequent comparative

analyses between B. oleracea linkage maps and the A. thaliana genome identified

numerous one-to-one segmental relationship and apparent genome duplication, in

addition to genome triplications (Lan et al. 2000; Babula et al. 2003; Lukens et al.

2003). Investigation of regions of the genome of B. oleracea containing specific genes

of interest also revealed various numbers of related genome segments (Quiros et al.

2001; Suzuki et al. 2003; Franzke et al. 2010). Recently, 186 miRNAs belonging to

55 families in B. rapa were identified by using comparative genomics (Dhandapani et

al. 2011).

Understanding the relative order of genome-wide duplication and taxonomic

divergence is central to comparative genomic biology (Kellogg 2003; De Bodt et al.

2005; Yu et al. 2011). Several studies have identified extensive local duplication in

the genome of Brassica species through physical and genetic mapping, and that the

genome has undergone triplication since their split from A. thaliana ca. 20 million

years ago (Mun et al. 2009; Beilstein et al. 2010; Proost et al. 2011). Comparison of

genome sequence of A. thaliana with BAC sequences from Brassica has revealed

conserved collinearity of gene order and content restricted to specific chromosomal

segments (Parkin et al. 2002; Qiu et al. 2009). Town et al. (2006) unravelled the gene

loss, fragmentation and dispersal after polyploidization in B. oleracea in context to A.

thaliana. In addition, comparison of pollen coat genes across Brassicaceae highlighted

rapid evolution by repeat expansion and diversification (Fiebig et al. 2004).

Apart from understanding genome and gene evolution, comparative genomics has

become a powerful tool in discovery of conserved non-coding sequences (Freeling

and Subramanium 2009; Meireless-Filho and Stark 2009) as in grasses (Guo and

Moose 2003) and miRNAs in Brassicaceae (Warthmann et al. 2008).

Genomic comparisons are also a quick way to prove suspect gene model annotations,

and to transfer genomic knowledge acquired in one taxon to a less studied one.


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Aligning syntenic regions of Arabidopsis chromosomes with their syntenic

chromosomal regions in other rosids allowed researchers to identify patterns of

conservation and divergence in the structure of genomes. The power and utility of

comparative genomics has led to the development of PLAZA, a web resource to study

gene and genome evolution in plants (Proost et al. 2009).

2.2. Regulation of Gene Expression in Plant Development

Growth and development of multi-cellular organisms is characterized by the

specification and differentiation of diverse cell types and organs which is controlled

by accurate spatio-temporal regulation of gene expression. Therefore, their abundance

needs to be tightly co-ordinated and controlled in both space and time, which is done

through precise regulation of gene expression. The process of gene regulation can

occur at various levels including at post-transcriptional level through control of

mRNA processing and stability, mRNA transport to the cytoplasm, translational, and

post translational modification (Baginsky et al. 2010). Small RNAs which are single

stranded RNA molecules of ~19-25 nucleotides (Bartel 2004) are one of the key

members of this regulatory network which plays important role in the process of gene

regulation at post-transcriptional level by targeting various transcription factors

(Vaucheret 2006). Based on their mode of action and biogenesis it has been

categorized as short interfering RNAs (siRNA), microRNA (miRNA), ta-siRNA and

piwi-interacting RNA (piRNA). Among these, microRNAs are a group of small

regulatory RNA which has generated considerable excitement in the field of various

developmental processes of plants and animals. As the present work deals with

characterization of miRNA gene and its regulatory mechanism, a comprehensive

account of related literature is provided.

2.3. MicroRNAs: An Overview

MicroRNAs, also called riboregulator are genomically encoded, evolutionarily

conserved, 20-24 nucleotides non-coding RNA which modulate gene expression in

plants and animals. It was first identified through forward genetics mapping of lin-4

mutant of Caenorhabditis elegans (Lee et al. 1993; Wightman et al. 1993). Analysis


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of this mutant showed that the lin-4 locus encodes a transcript that could give rise to a

fold-back structure and small 21 nt RNA fragment. This lin-4 encoded small RNA

could suppress the expression of target Lin-14 and Lin-28 by pairing with the 3’UTR

(Lee et al. 1993; Wightman et al. 1993). Subsequently, another miRNA, let-7 was

identified from C. elegans that targets in similar manner and silences lin-41 (Bagga et

al. 2005). Among plants, a total of 2985 microRNAs have been reported from a wide

variety of species (www.mirbase.org, release 18.0, November 2011). Such as,

Arabidopsis thaliana, Brassica napus, Glycine max, Medicago truncatula, Oryza

sativa, Physcomitrella patens, Populus trichocarpa, Saccharum officinarum, Sorghum

bicolor, Zea mays, Chlamydomonas reinhardtii, Selaginella moellendorffii, Malus

domestica, Populus euphratica and Pinus taeda (Table 2.1) and also other. These

miRNA can be categorized into 1093 families (Reinhart and Bartel 2002; Rhoades

and Bartel 2004; Wang et al. 2004; Billoud et al. 2005; Zhang et al. 2005).


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Table 2.1: An overview of some miRNAs and families found in plants (as per miRBase 18.0 release 2011, www.mirbase.org)

Plants Total microRNA Families Precursors Mature

Eudicotyledons

Arabidopsis thaliana 291 328 195 Arabidopsis lyrata 201 375 121

Brassica napus 46 48 17 Brassica oleracea 6 7 5

Brassica rapa 19 23 10 Carica papaya 1 1 1

Arachis hypogaea 23 32 21 Glycine max 362 395 120 Glycine soja 13 13 9

Lotus japonicus 3 4 3 Medicago truncatula 335 674 126 Phaseolus vulgaris 8 10 8

Vigna unguiculata 18 18 1 Rehmannia glutinosa 6 6 6 Gossypium arboretum 1 1 1 Gossypium herbacium 1 1 1 Gossypium hirsutum 34 36 19 Gossypium raimondii 4 4 3

Theobroma cacao 82 82 24 Aquilegia coerulea 45 45 20

Malus domestica 1 2 1 Citrus clementine 5 5 4 Citrus reticulata 4 4 3

Citrus sinensis 60 64 40 Citrus trifoliata 6 6 6

Populus euphratica 5 5 4 Populus trichocarpa 234 237 40

Ricinus communis 63 63 22 Solanum lycopersicum 37 37 23

Vitis vinifera 163 186 49 Monocotyledons

Aegilops tauschii 2 2 2

Brachypodium distachyon 142 143 76 Festuca arundinacea 15 15 13

Hordeum vulgare 22 23 18 Oryza sativa 581 661 206

Saccharum officinarum 16 16 6 Saccharum spp. 18 20 13 Sorghum bicolor 171 172 27 Triticum aestivum 44 44 39 Triticum turgidum 1 1 1

Zea mays 172 321 28 Chlorophyta Chlamydomonas reinhardtii 50 85 47

Picea abies 40 41 32 Coniferophyta Pinus taeda 37 38 26 Embryophyta Physcomitrella patens 229 280 108

Selaginella moellendorffii 58 64 45


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2.3.1. Evolution of miRNA genes in plants

In the past few years the origins of miRNA genes have attracted extensive attention,

and a variety of hypotheses have been continuously proposed (Allen et al. 2004;

Smalheiser and Torvik 2006; Maher et al. 2006; Piriyapongsa et al. 2007;

Piriyapongsa and Jordan 2008; Guo et al. 2009; Zhang et al. 2011a). There are very

few common miRNAs that have been discovered in plants and animals (Lu et al.

2005; Arteaga-Vázquez et al. 2006) which might suggest independent evolution of

miRNAs. Plant miRNAs probably originated from inverted duplication of target gene

sequences, followed by accumulation of mutations (Jasinski et al. 2010; Ma et al.

2010; Fahlgren et al. 2010; Axtell et al. 2011; Zhang et al. 2011a; Figure 2.1).

Sequence divergence at the inverted duplication locus occurred under constraints to

maintain both the fold-back structure and recognition by DICER like 1 (DCL1).

Sequence degeneration continued until the point that only the miRNA or miRNA

complementary sequences were maintained for matching the founder gene sequence

(Allen et al. 2004; Axtell et al. 2011). Presence of non-conserved and potentially

evolutionary young miRNAs located adjacent to their respective target genes

containing sequence similarities to these genes also outside the small RNA sequence

can be taken as evidence of evolution of miRNAs (Allen et al. 2004; Olena and Patton

2010). The systematic analysis of miRNAs genes between A. thaliana and A. lyrata

showed that there is sequence divergence in stem-loop region while miRNA region is

consistent due to requirement for complementarity to target genes and thus, remains

evolutionary conserved in function. In contrast, the miRNA genes which are more

divergent even in miRNA region because of the natural evolution of younger genes

caused the loss of target complementarity and therefore would not be expected as

functional miRNA (Ma et al. 2010; Fahlgren et al. 2010).


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Figure 2.1: Evolution of miRNA through inverted gene duplication

2.3.2. Biogenesis and mechanism of microRNAs

MicroRNAs are encoded in the genome as independent transcription units (Lee et al.

2002a; Cui et al. 2009) and transcribed by RNA polymerase II (RNA pol II) (Lee et

al. 2004; Cui et al. 2009) and further processed in association with various

components (Table 2.2; Figure 2.2). The presence of CpG island, TATA box,

initiation elements and histone modifications in miRNA gene shows that the promoter

region is similar to the protein coding genes (Ozsolak et al. 2008; Corcoran et al.

2009) and are controlled by transcription factors, enhancers and via chromatin

modification. Recent research demonstrates that the transcription of miRNA also

occurs through RNA pol III (Borchert et al. 2006; Diebel et al. 2010). RNA

polymerase II (RNA Pol II) generates primary miRNA (pri-miRNA) with 5’ capped

and 3’ polyadenylated structure, ranging in size from hundreds of nucleotides to tens

of kilo bases (Cai et al. 2004; Lee et al. 2004). The primary to precursor- miRNA

(pre-miRNA) conversion and release of the mature miRNA are catalyzed by DCL1 in

plants. The conversion of pri-to-pre-miRNA also necessitates the double-stranded


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RNA-binding protein HYPONASTIC LEAVES1 (HYL1) and the C2H2-zinc finger

protein SERRATE (SE), which interact with DCL1 in nuclear processing centers

called D-bodies or SmD3/SmB-bodies (Kurihara et al. 2006; Fang and Spector 2007).

HYL1 belongs to a family of dsRNA-binding proteins in Arabidopsis (Lu and

Fedoroff 2000; Hiraguri et al. 2005; Wu et al. 2007). SE encodes a C2H2 zinc finger

protein that was initially found to specify leaf polarity through promoting the

accumulation of miR165/166 (Grigg et al. 2005). Later, it was demonstrated that SE

plays a general role in the biogenesis of many miRNAs (Lobbes et al. 2006; Yang et

al. 2006). In plants, DCL1 cleaves pre-miRNA in the nucleus but in animals, the pre-

miRNA is first exported from the nucleus, and the canonical Dicer enzyme carries out

the cleavage reaction in the cytoplasm (Kim 2005). The final miRNA: miRNA*

duplex are characterized by the presence of 2 nucleotide overhangs at 3’ end and free

5’ phosphate, a characteristic feature of DCL1 products (Elbashir et al. 2001;

Havecker et al. 2010). MicroRNAs show different characteristics between plants and

animals. In animals miRNA: miRNA* is processed by Drosha, RNase III enzyme in

association with Pasha (Lee et al. 2003). Export of duplex from nucleus to cytoplasm

is accomplished by HASTY (Telfer and Poething 1998), the orthologs of Exportin-5

in animals, in co-operation with RAN1-GTP (Bollman et al. 2003). Accumulation of

some miRNAs is not affected in hasty mutants, suggesting the existence of HASTY-

independent miRNA export systems for example, in association with AGO1

(Argonaute 1) protein (Farazi et al. 2008). Mature miRNA duplexes are stabilized by

the S-adenosyl methionine-dependent methyltransferase-HUA ENHANCER1

(HEN1), which methylates small RNAs (Yu et al. 2005b). Methyl groups added to the

3’ terminal nucleotides of each strand prevent their uridylation and subsequent

degradation. The strand with lower thermodynamic stability at its 5’ end is

subsequently incorporated into RNA-induced silencing complex (RISC) and interact

with AGO protein member to regulate gene expression at transcriptional, post-

transcriptional and translational level (Khvorova et al. 2003; Wypijewski et al. 2009).

AGO protein family is an integral part of the ribonucleoprotein complex called RISC

(Bartel 2004; Farazi et al. 2008; Höck and Meister 2008; Thomson and Lin 2009).

These effector complexes mediate different small RNA functions at the transcriptional


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and/or post-transcriptional level, such as target mRNA cleavage, translational

repression, and regulation of chromatin structure. Different members of the AGO

family often show distinct tissue distribution, which allows them to mediate tissue-

specific small RNA functions (Havecker et al. 2010).

Table 2.2: Factors involved in small RNA biogenesis

Name Type Substrate Activity

Argonaute (AGO)

RNase H endonuclease

Precursor miRNAs Component of RISC complex, cleaves passenger strand of some miRNA precursors or act as RNA slicer

Dicer Like Protein (DCL)

RNase III endonuclease

Precursor miRNAs Generates hairpin precursor, removes loop from precursor to generate mature miRNA duplex.

Hua Enhancer1 (HEN1)

Methyltransferase Mature miRNAs and siRNAs in plants;

piRNAs and siRNAs in animals

Adds 2′-O-methyl group to the 3′ ends of small RNAs for stabilization and protection from degradation by SDN1.

Hyponastic Leaves1 (HYL1)

dsRNA-binding protein

Primary miRNA transcript

Stabilization of pri-miRNA

HASTY Exportin-5 ortholog

Mature miRNAs Transport of miRNA:miRNA* from nucleus to cytoplasm

Small RNA Degrading Nuclease (SDN1)

3′-to-5′ exonuclease

Mature single-stranded miRNAs

Degrades mature miRNAs

DAWDLE (DDL)

RNA-binding protein

RNA stabilizer Stabilize the stemloop for adaptation into D-bodies and help Dicer-like-1 (DCL1) precursor recognition

SERRATE (SE) C2H2-zinc finger double stranded RNA binding

protein

Primary miRNA transcript

Stabilization of pri-miRNA


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Figure 2.2: Biogenesis of miRNAs in plant The miRNA gene is transcribed by RNA polymerase II (pol II) to generate primary transcript (pri-miRNA). The RNA-binding protein DAWDLE (DDL) stabilizes the stem-loop for adaptation into D-bodies (nuclear processing centre) and also interacts with Dicer-like-1 (DCL1) to help stem-loop precursor recognition. In this process, rigorous interaction of C2H2-zinc finger protein SERRATE (SE), the double stranded RNA binding protein HYPONASTIC LEAVES1 (HYL1) and nuclear cap binding protein (CBC) play important role. The mature miRNA duplex expurgated from the pri-miRNA (miRNA/miRNA*; where miRNA in black is guide strand and miRNA* in blue is degraded strand) and the terminal methyl group is added by the S-adenosyl methionine (SAM)-dependent methyltranferase-HUA ENHANCER1 (HEN1) to prevents the duplex from being degraded by the small RNA degrading nuclease (SDN). HASTY export the miRNA/miRNA* from the nucleus to the cytoplasm. The guide miRNA strand is incorporated into RNA induced silencing complex (RISC) to carry out target mRNA cleavage or translational repression.


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2.3.3. MicroRNAs as regulator of developmental pathways in plants

It is amply evident that gene regulation is mediated through the action of microRNAs

that play critical role at each major stage of plant and animal development (Rhoades

et al. 2006). A good number of miRNAs and their target genes have been identified

and demonstrated to regulate various plant developmental processes including

flowering time control, floral development, leaf polarity and morphogenesis and root

development (Palatnik et al. 2003; Aukerman and Sakai 2003; Emery et al. 2003;

Chen 2004; Juarez et al. 2004; Laufs et al. 2004; Mallory et al. 2004a,b; McHale and

Koning 2004; Vaucheret et al. 2004; Kidner and Martienssen 2005a; Meng et al.

2010; Song et al. 2011). MicroRNAs not only regulate transcription factors, but also

modulate their own biogenesis and function and therefore, they typically act as a core

component of gene regulatory network and implicated as an important factor in plant

development (Rhoades et al. 2002, Mallory et al. 2004a; Vaucheret 2006; Table 2.3).

In addition, studies have also demonstrated that small RNAs including miRNAs are

mobile and can be transported through the phloem system from one organ to another

(Chitwood and Timmermans 2010). Therefore, miRNAs could be involved in long-

distance signal transduction (Buhtz et al. 2008; Pant et al. 2008; Kanehira et al. 2010).

The role of miRNAs in the regulation of plant development is supported by various

developmental phenotypes due to loss-of-function alleles of genes involved in their

biogenesis and function. Hypomorphic alleles of DCL1 reduce the production of most

miRNAs (Park et al. 2002) and display a range of developmental phenotypes,

including defects in leaf morphology, axillary meristem maintenance, flowering time,

inflorescence determinancy, floral organ patterning and ovule development, while

null alleles are embryonic lethal (Jacobsen et al. 1999; Schauer et al. 2002; Golden et

al. 2002). Mutations in AGO1 cause unregulated or ectopic expression of genes

targeted by miRNAs and also causes a decrease in the abundance of some miRNAs

(Vaucheret et al. 2004; Kidner and Martienssen 2005b; Ronemus et al. 2006). AGO1

hypomorphic alleles have defects in lateral organ polarity, leaf and flower

morphology (Vaucheret et al. 2004; Kidner and Martienssen 2005b). Studies in A.

thaliana and rice have demonstrated that members of the Argonaute (AGO) family


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are essential regulators of SAM (Nagasaki et al. 2007; Nogueira et al. 2007; Liu et al.

2009). More specifically, it has been shown that AGO10 represses miRNA165/166

(miR165/166) for proper SAM-maintenance as well as the establishment of leaf

polarity (Liu et al. 2009; Zhu et al. 2011). Wong et al. (2011) provides additional

information towards understanding the role of key regulators in regulatory circuits in

the SAM of soybean during shoot development.

Apart from the role of miRNAs in development it is also involved in regulation of

adaptive traits such as abiotic stress. Several studies demonstrate that drought and

salinity stresses induce differential expression of miRNAs in a variety of plant

species, including Arabidopsis (Liu et al. 2008; Jagadeeswaran et al. 2009; Jia et al.

2009), rice (Zhao et al. 2007a), maize (Ding et al. 2009), Poplars (Lu et al. 2005; Jia

et al. 2009), and tobacco (Frazier et al. 2010; Frazier et al. 2011). Currently, a number

of miRNAs-miR156, miR159, miR165, miR167, miR168, miR169, miR319, miR393,

miR395, miR396, miR398, miR399, and miR402 have been reported to be induced by

drought and salinity stress in several plant species. Over-expression of miR169

confers enhanced drought tolerance in tomato (Zhang et al. 2011b).


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Figure 2.3: Regulatory network of miRNAs and its targets involved in various

developmental stages of plant (adapted from Wang et al. 2007)


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Table 2.3: Examples of some miRNAs involved in plant development (Modified and updated from Wang et al. 2007)

Development events miRNAs miRNA targets References Leaf development, Patterning and polarity

miR165/166

miR164a

miR319/JAW

miR159

HD-ZIP III transcription factor

NAC-containing TF: CUC2

TCP2, TCP3, TCP4, TCP10, TCP24 MYB TFs: MYB33, MYB65

Zhong and Ye 2004; Zhou et al. 2007; Pulido et al. 2010 Nikovics et al. 2006; Kim et al. 2009. Palatnik et al. 2003. Palatnik et al. 2003; Millar and Gubler, 2005.

Floral identity and flower development

miR172

miR164c

miR159

APETALA2-like TFs: AP2, TOE1, TOE2, TOE3

CUC1, CUC2 MYB TFs: GAMYB, MYB33, MYB65

Chen 2004; Schwab et al. 2005; Mlotshwa et al. 2006; Zhao et al. 2007b. Baker et al. 2005. Achard et al. 2004; Millar and Gubler 2005; Schwab et al. 2005; Tsuji et al. 2006.

Flowering time miR159

miR172

miR156

miR171

MYB TFs: GAMYB

AP2, TOE1, TOE2, TOE3

SBP-like TFs: SPL3 SCL TFs: SCL6-II, SCL6-III, SCL-IV

Achard et al. 2004; Schwab et al. 2005. Mlotshwa et al. 2006; Yant et al. 2010. Schwab et al. 2005; Wang et al. 2009. Llave et al. 2002.

Developmental phase transition

miR172

miR156

APETALA2-like TFs: GL15 SBP-like TFs: SPL3, SPL4, SPL5

Aukerman and Sakai 2003; Lauter et al. 2005. Schwab et al. 2005; Luo et al. 2006; Xing et al. 2010

Shoot and root development

miR164

miR160

NAC-containing TF: CUC1, CUC2, NAC1 ARF10, ARF16, ARF17

Rhoades et al. 2002; Laufs et al. 2004; Schwab et al. 2005; Guo et al. 2005. Meng et al. 2010.

Vascular and plastid development

miR166 HD-ZIP TFs: ATHB15 Kim et al. 2005; Williams et al. 2005; Ochando et al. 2006; Donner et al. 2009.

Hormone signalling for plant development

miR159

miR160

miR167

miR164

miR393

MYB TFs: GAMYB

ARF TFs: AFR10, AFR16, AFR17 ARF TFs: AFR6, AFR8

NAC-containing TF: NAC1

F-box protein: TIR1

Achard et al. 2004; Schwab et al. 2005. Mallory et al. 2005; Wang et al. 2005. Rhoades et al. 2002; Ru et al. 2006; Wu et al. 2006. Guo et al. 2005. Rhoades and Bartel 2004; Sunkar and Zhu 2004; Si-Ammou et al. 2011.


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2.3.3.1. Hormone signaling

Plant hormones such as auxin, gibberellin and absicic acid (ABA) play significant

roles in developmental processes such as embryogenesis, cell division, elongation,

differentiation and organ formation. A number of miRNAs such as miR159, miR160,

miR164, miR167 and miR393 have been reported to respond to such plant hormones

(Sunkar and Zhu 2004; Zhang et al. 2005). As auxins are involved in almost all

aspects of plant development (Wang et al. 2005), inexplicably, a number of genes are

reported which are involved in auxin signalling. Among those, few are known to act

as targets of miRNAs such as auxin response elements (AREs) (Rhoades et al. 2002;

Rhoades and Bartel 2004; Sunkar and Zhu 2004; Mallory et al. 2005;). The transport

inhibitor response 1 (TIR1) auxin receptor is a predicted target of miR393 (Bonnet et

al. 2004; Wang et al. 2004; Adai et al. 2005). A recent report states that in

Arabidopsis, miR393 regulates auxin signaling and auxin-mediated functions by

down-regulating the expression of TIR1/AFB2 Auxin Receptor (TAAR) genes (Si-

Ammou et al. 2011). In addition, miR164 is induced by auxin to clear NAC1 mRNA

to reset auxin signals (Guo et al. 2005). Furthermore, ARF10, ARF16 and ARF17 are

predicted targets of miR160, and ARF6 and ARF8 are targets of miR167. Disruption of

miR160 results in increased levels of expression of ARF10, ARF16 and ARF17 and

distorted expression of auxin early responsive genes which consequently cause severe

developmental defects (Mallory et al. 2005; Gutierrez et al. 2009). Auxins play a key

role in setting up the position of leaf primordia and allowing outgrowth. In very

young organ primordia expression of the PIN1 auxin efflux carrier marks the abaxial

boundary of REVOLUTA (REV) expression (Heisler et al. 2005). Several reports have

described the regulation of leaf polarity by the miR390/ARF pathway (Nagasaki et al.

2007; Chitwood et al. 2009; Schwab et al. 2009). On the other hand, gibberellic acid

(GA) has been shown to modulate miR159 levels during anther development (Achard

et al. 2004).

2.3.3.2. Flower development

Flower development is one of the most important stages of plant development. The

regulation of floral organ development is carried out by the combinatorial action of


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genes explained by the ABC/ABCDE model (Lohmann and Weigel 2002; Jack 2004;

Alvarez-Buylla et al. 2010; Causier et al. 2010; Ito et al. 2011). The combination and

communication of different classes of transcription factors connect to recreate floral

architecture and flowering time. However, some members of these gene classes are

regulated by miRNAs (Chen 2004). APETELA2 (AP2), a class A gene, is a target of

miR172 and overexpression of miR172 in Arabidopsis disrupts the specification of

floral organ identity and causes early flowering (Aukerman and Sakai 2003) while

overexpression of an AP2 results in late flowering (Yant et al. 2010). Therefore,

miR172 plays a major role in regulating floral patterning, identity, and flowering time

(Chen 2004; Zhao et al. 2007b; Zhu and Helliwell 2011). MicroRNA resistant forms

of AP2 cause an enlarged and undefined floral meristem, floral organ patterning

defects, and extra whorls of stamens (Chen 2004). Zhao et al. (2007b) have shown a

complex pathway for AP2 gene which acts through the WUSCHEL and AGAMOUS

pathways to affect floral meristem size, determinacy, and organ identity (Zhao et al.

2007b). Studies revealed that miR172 regulates flowering time through the repression

of the AP2, TOE1 and TOE2 (Aukerman and Sakai 2003). The involvement of

miR172 in sex determination and meristem branching in maize inflorescence was

discovered by the cloning of two classic maize mutants, the recessive tasselseed4

(ts4) mutant and the dominant Tasselseed6 (Ts6) mutant (Irish 1996). Studies in

maize, rice, and barley exibit that miR172 is also important in regulating phase

transition and determination of floral organ identity in monocotyledons (Lauter et al.

2005; Chuck et al. 2007a; Zhu et al. 2009; Nair et al. 2010). Conversely, Xing et al.

(2010) reported that development of fully fertile flowers in Arabidopsis require the

action of multiple miR156/7-targeted SPL genes in concert with SPL8.

2.3.3.3. Leaf development

Leaves are vital organs of plants and its normal shape and size serves as an important

factor that influences the proper growth and development through photosynthesis. To

date, at least five miRNAs (miR156, miR159, miR165, miR166, and miR319) have

been identified which control the patterning and development of leaves in

Arabidopsis, maize, and other plant species (Pant et al. 2008; Jung et al. 2009;

Kanehira et al. 2010). These miRNAs regulate leaf development by targeting the HD-


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ZIP III and the TCP transcription factor genes. MiR319 is known to be involved in

proper development of leaf shape and curvature through regulating TCP transcription

factor genes (Palatnik et al. 2003). In addition to the conserved miRNAs, non-

conserved miRNAs may also play roles in leaf development. For instance, miR824 is

reported to play a role in stomatal development (Kutter et al. 2007). Furthermore,

overexpression of miR156, significantly results in increases leaf initiation and plant

biomass in Arabidopsis (Schwab et al. 2005). This suggests that miRNA-based

biotechnology can be employed for improvement of plant biomass for agriculture and

for biofuel production.

Basically, leaves originate from shoot apical meristem (SAM), a group of stem cells

at the apex of the shoot (Steeves and Sussex 1989). Patterning of leaf primordia into

an upper (adaxial) and lower (abaxial) side of the leaf is the key stage (Husbands et al.

2009) and is regulated by a complex regulatory network (Table 2.4). It has been

known that a signal from the meristem is required for the specification of the adaxial

side of the leaf (Sussex 1955). Heisler et al. (2005) and Reinhardt et al. (2005)

reported that this signal travels via epidermal layer. The identity of this signal

however, is currently unknown. Adaxial side of the leaf is exposed to the sun and

specialized for light capture with tightly packed columellar palisade cells and the

abaxial is specialized for transpiration with many stomata and loosely packed spongy

mesophyll cells. The specification of adaxial and abaxial sides of the leaf is vital for

generating the leaf blade or lamina which grows at the boundary between the two

sides of the leaf. Polarity of leaves is established through the antagonistic interactions

between genes of the transcription factor class III homeodomain leucine zipper (HD-

ZIP III) members i.e., PHAVOLUTA (PHV), PHABULOSA (PHB) and REVOLUTA

(REV) are regulated by miR165/166 and play significant role in regulation of leaf

polarity. Higher accumulation of miR165/166 results in altered adaxial and abaxial

identity (Williams et al. 2005; Kidner 2010; Rubio-Somoza and Weigel 2011).

Dominant mutations in the member of HD-ZIP III members resulted in abnormal

adaxialized and radicalized leaf patterning (McConnell et al. 2001; Emery et al. 2003;

Zhou et al. 2007). Correct accumulation of miR165/166 is influenced by the

expression pattern of AGO1 and PNH/ZLL (Kidner and Martienssen 2004; Liu et al.


22

2009). Recently Zhu et al. (2011) reported that AGO10 has a unique property to bind

miR165⁄166 preferentially over other miRNAs. They also showed that AGO10

competes with AGO1, the predominant component of the RISC in tissues, to

specifically bind miR165⁄166 and attenuate its regular action to repress target HD-ZIP

III proteins. HYPONASTIC LEAVES1 (HYL1) is another important regulator to

monitor the role of miR165/166, miR319a, and miR160 in leaf flattening through the

relative activities of adaxial and abaxial identity responsive genes and thus playing an

essential role in leaf development (Dong et al. 2008; Liu et al. 2011). Recent findings

suggest that the Dof5.1 protein directly binds to the REV promoter and regulates

adaxial-abaxial polarity of leaf (Kim et al. 2010).

Studies have revealed that miR164 is another regulator of leaf patterning. The balance

between miR164a and the transcription factor, CUP-SHAPED COTYLEDON 2

(CUC2) controls leaf margin serration in Arabidopsis. Although the pattern of

serration is determined first independently of CUC2 and miR164, the balance between

co-expressed CUC2 and miR164a determines the extent of leaf serration (Nikovics et

al. 2006). As with HD-ZIP III, TCP and the miR319/JAW family are found in a wide

range of plant species, suggesting that miRNA-mediated control of leaf

morphogenesis is conserved across different plant species with different leaf forms

(Palatnik et al. 2003). MiR390–TAS3–ARF has been invoked in the control of leaf

margin growth in both Arabidopsis and Lotus japonicus (Fahlgren et al. 2006; Yan et

al. 2010). According to previous reports, overexpression of miRNA-resistant version

of MYB33 in Arabidopsis resulted in upward curling of leaves (Palatnik et al. 2003).

Similarly, Arabidopsis transformed with MYB33 containing the mutated miRNA

target site show dramatic pleiotropic developmental defects, including abnormal

leaves that were rounded and upturned at the sides (Millar and Gubler 2005).


23

Table 2.4: Some examples of determinants of adaxial and abaxial fate (adapted

from Kidner and Martienssen 2004; Kim et al. 2010)

Protein or small RNA

family

Expression in leaf primordia

Functions in Arabidopsis

Regulation

miR165/166 Abaxial Abaxial determinant, vascular patterning

HD-ZIPIII transcripts regulation

HD-ZIP III Adaxial Adaxial determinant, vascular patterning, meristem function

Repressed on abaxial side by miR166 and KANADI proteins

ARP Uniform Adaxial determinant, KNOX repression

Acts together with AS2, redundantly to the ta-siRNA pathway, and in opposition to ETT-ARF4

AS2 Unknown Adaxial determinant, KNOX repression

Acts together with AS1, redundantly to the ta-siRNA pathway, and in opposition to ETT-ARF4

KANADI Abaxial Abaxial determinant, vascular patterning

Repressed on adaxial side by HD-ZIP III proteins

ARF ETT/ARF3: uniform; ARF4:

abaxial

Abaxial determinant, auxin signaling

Targets of TAS3 ta-siRNAs, act in opposition to AS1-AS2

YABBY Abaxial Abaxial determinant, blade outgrowth

Act downstream of all other known polarity determinants

TAS3 ta-siRNA

Unknown Unknown Cleaves ETT and ARF4 transcripts

Dof5.1 protein

Unknown adaxial-abaxial regulator Binds to the REV promoter

2.3.3.4. Vascular development

Vascular system is an intricate network of conducting tissues that interconnects all

plant organs and transports water, minerals, organic compounds, and signalling

molecules throughout the plant body. It consists of two conducting tissues, xylem and


24

phloem and procambial/ cambial cells (Steeves and Sussex 1989). Leaf development,

its shape and functions are dependent on its vasculature which not only provides

mechanical support, but also supply metabolites and signalling molecules (Scarpella

et al. 2010). It is now accepted that a unified molecular mechanism modulates

temporal and spatial development of vascular tissues in different plant species,

although vascular pattern and organization are quite diverse (Baima et al. 2001).

MicroRNA165/miR166 and miR159 have indispensable roles in the development of

leaf vasculature. Regulation of ATHB8 through miR165/miR166 allows progression of

vasculature differentiation (Allen et al. 2007; Donner et al. 2009; Alonso-Peral et al.

2010), although, miR166 mediated regulation of ATHB15 through mRNA cleavage is

a principal mechanism for the regulation of vascular development. Gain-of-function

miR166a mutant results in decreased transcript level of ATHB15 that was

accompanied by an altered vascular system with expanded xylem tissue and

interfascicular region and inactivation of vascular cell differentiation from

cambial/procambial cells (Kim et al. 2005). Apart from these, growth hormones like

auxins and brassinosteroids also play regulatory role in vascular tissue differentiation

(Jang et al. 2000; Sachs 2000; Carland et al. 2002). The potential interplay between

miRNA and its target in leaf vasculature development based on contributions of AS1

and miR159-targets to downregulation of miR165/miR166 expression is yet to be

investigated in detail (Li et al. 2005; Yang et al. 2008b).

2.4. Transcription Factors

Transcription factors (TFs) are essential for regulation of gene expression. The effect of TFs on gene expression is a significant event and the consequence of TF-mediated gene regulation in a developmental context can be usually classified as a switch that determines the precise composition of a gene network and therefore defines the execution of a cellular differentiation program. Interaction of TFs with combinations of cis-motifs within promoter region in a sequence specific manner accounts for the specificity of gene expression (Singh 1998). Sequences towards 3′ of the coding region are essential for transcription termination and polyadenylation. In some cases, regulatory elements are also present within introns or the gene coding region (Bolle et al. 1996; Schauer et al. 2009). Developmental and tissue specific gene expression is regulated by the interaction with enhancers and promoters with general and tissue specific DNA binding factors (Bondino et al. 2009; Siefers et al. 2009). Based on the


25

DNA binding motifs, TFs in plants can be classified into different groups that are involved in control of morphogenesis, differentiation and adaptation. These domains are the most conserved regions within genes of the same family. Table 2.5 provides a partial list of some of the major families of DNA-binding domains/transcription factors.

Table 2.5: Structural features of some conserved domains used to classify plant

transcription factors (Liu et al. 1999)

S. No. DNA-binding Domain Domain Architecture

1 Homeodomain (HD) Approximately 60 amino acid residues producing either three or four α-helices and an N-terminal arm

2 Zinc finger (ZF) Finger motif (s) each maintained by cysteine and/or histidine residues organized around a zinc ion

3 bZIP A basic region and a leucine-rich zipper-like motif 4 Myb- related A basic region with one to three imperfect repeats each

forming a helix-helix-turn-helix 5 Trihelix Basic, acidic and proline/glutamine-rich motif which

forms a trihelix DNA-binding domain 6 Basic helix-loop-helix

(bHLH/Myc) A cluster of basic amino acid residues adjacent to a helix-loop-helix motif

7 MADS Approximately 57 amino acid residues that comprise a long α-helix and two β-strands

8 AT- hook motif A consensus core sequence R(G/P)RGRP with the RGR region contacting the minor groove of A/T-rich DNA

9 HMG-box L-shaped domain consisting of three α -helices with an angle of about 80o between the arms

10 APETALA2/ Ethylene Responsive Element Binding Protein (AP2/ EREBP)

A 68-amino acid region with a conserved domain that constitutes a putative amphiphatic α-helix

11 B3 A 120 amino acid conserved sequence at the C-termini of VP1 and ABI3

12 Auxin Responsive Factor (ARF)

A 350 amino acid region similar to B3 in sequence

13 MYB domain A basic region with one to three 50-53 amino acids imperfect repeats that form the helix-turn-helix motif.

14 NAC domain A twisted β-sheet surrounded by a few helical elements. 15 WRKY domain A domain constituted by around 60 amino acids with a

conserved WRKYGQK sequence is followed by a C2H2- or C2HC-type of zinc finger motif.


26

2.4.1. HD-ZIP transcription factor family

The HD-ZIP family of transcription factors are unique to the plant kingdom and is

comprised of by more than 25 genes in A. thaliana (Schena and Davis 1992; Elhiti

and Stasolla 2009). Special features of HD-ZIP members are the presence of a

homeodomain (HD) and a leucine zipper motif (ZIP). A combination of

homeodomain and leucine zipper proteins acts as a dimerization motif (Lee and Chun

1998). On the basis of their distinguishing characteristics HD-ZIP family have been

classified into four subfamilies: (i) conservation of the HD-ZIP domain that

determines DNA-binding specificities, (ii) genes structures, (iii) additional conserved

motifs and (iv) functions (Table 2.6; Figure 2.4).

Table 2.6: Classification of HD-ZIP transcription factors on the basis of their

functions

Subfamily Functions References

HD-ZIP I Response to abiotic stress, ABA, de-

etiolation, blue-light signaling

Wang et al. 2003; Olsson et al.

2004; Henriksson et al. 2005.

HD-ZIP II Response to illumination conditions,

shade avoidance, auxins

Sessa et al. 1998; Sawa et al.

2002; Rueda et al. 2005.

HD-ZIP III Embryogenesis, meristem regulation,

lateral organs initiation, leaf polarity,

vascular system development, auxin

transport

McConnell et al. 2001; Mattsson

et al. 2003; Prigge et al. 2005;

Williams et al. 2005; Emery et al.

2003; Kim et al. 2005.

HD-ZIP IV Epidermal cells differentiation,

anthocyanin accumulation, root

development, trichomes formation

Kubo et al. 1999; Luo and

Oppenheimer 1999; Nakamura et

al. 2006.


27

Figure 2.4: Schematic representation of HD-ZIP sub-families exhibiting distinctive features Abbreviations: CPSCE- named after the five conserved amino acids Cys (C), Pro (P), Ser (S), Cys (C), Glu (E), adjacent to and downstream of LZ, and an N-terminal consensus sequence; MEKHLA domain- named after the highly conserved amino acids Met (M), Glu (E), Lys (K), His (H), Leu (L), Ala (A); N-Term-N-terminus consensus; START-steroidogenic acute regulatory protein-related lipid transfer domain; HD- Homeodomain; LZ- Leucine Zipper; SAD- Smad 4 activation domain is a proline-rich, p300-dependent transcriptional activation domain. HD-ZIP-encoding genes have been isolated from a wide variety of plants, such as

Solanum lycopersicum, Craterostigma plantagineum, Zea mays, Pisum sativum,

Glycine max, Daucus carota, Heliantus annuus, Nicotiana sylvestris, Silene latifolia,

Picea excelsa, Zinnia elegans, Lotus japonicus, Medicago truncatula, Brassica napus

and Physcomitrella patens which includes monocots and dicots, C3 and C4 plants

(Valle et al. 1997; Sakakibara et al. 2001; Deng et al. 2002; Ingouff et al. 2003; Ageez

et a., 2003; Tron et al. 2004; Rueda et al. 2005; Yu et al. 2005c; Manavella et al.

2006).

On the basis of evolutionary studies HD-ZIP III genes show remarkable conservation

with LZ motif and START domain-containing proteins that were first classified in this

sub-family only (Schrick et al. 2004). Phylogenetic analysis revealed that HD-ZIP III

genes were the first which associated with basic growth and patterning in ancient land

plants. Throughout evolution, they diversified and acquired new functions that

contributed to the modification of land plant development and to the origin of new


28

tissues and organs, such as the vascular system and leaves (Prigge and Clark 2006;

Floyd et al. 2006).

2.5. Conjugative Role of miRNAs and Transcription Factors

in Plant Development

Developmental programs involve multilayered TFs network, cascades and regulatory

loops (Davidson 2001); in other words, miRNA expression patterns are determined by

complex transcriptional regulatory inputs. Transcription factors (TFs) and

microRNAs (miRNAs) are the largest families of trans-acting gene regulatory

molecules in multicellular organisms and they act in a largely combinatorial manner

(Table 2.7). There are several lines and evidences which support the regulation of TFs

by miRNAs. For example, hierarchial action of miR156, miR172 and their targets SPL

and AP2 control the vegetative phase change and flowering in the annual grasses, rice

and maize (Poethig 1988; Evans et al. 1994; Moose 1994; Lauter et al. 2005; Xie et

al. 2006; Chuck et al. 2007b; Salvi et al. 2007). In addition, miR164 targets the NAC1

(CUC1) domain encoding mRNAs which transduces auxin signals for lateral root

emergence. Inducible expression of miR164 by auxin in wild-type plants led to

decreased NAC1 mRNA levels and reduced lateral root emergence (Guo et al. 2005).

ARF10, ARF16 and ARF17 targeted by miR160 and the mechanism of conjugative

action of ARF10 and miR160 have been analyzed using de-repression experiments

through creating silent mutations in the ARF10 sequence complementary to miR160

(Liu et al. 2007). Palatnik et al. (2003) reported that miRNA resistant version of

MYB33 plants developed curled-up leaves indicating that the down regulation of

MYB33 is important for normal leaf development. Millar and Gubler (2005) also

showed that transgenic plants containing a miR159-resistant version of MYB33

expressed under the control of its own promoter developed various phenotypes,

including leaf in curvature, rounded leaves with short petioles, reduced apical

dominance and low fertility. Thus, miRNA-target nodes play a pivotal role in

governing plastic behaviour during development such as phase change and plant

architecture and in response to the biotic and abiotic stresses (Rubio-Somoza and

Weigel 2011; Wang et al. 2011d; Khraiwesh et al. 2011). Furthermore, evolutionarily

well-conserved miRNAs are likely to contribute to proper plant growth and


29

morphogenesis by regulating their target (Axtell and Bowman 2008; Todesco et al.

2010).

Table 2.7: Some examples of targets of A. thaliana, O. sativa and P. trichocarpa

miRNAs (Adapted from Rhoades et al. 2006)

miRNA family Target transcription

factor family

Genes

miR165/166 HD-ZIPIII PHB, PHV, REV, ATHB-8, ATHB-15

miR156 SBP SPL2, SPL3, SPL4, SPL10

miR159 MYB MYB33, MYB65

miR319 TCP TCP2, TCP3, TCP4, TCP10, TCP24

miR160 ARF ARF10, ARF16, ARF17

miR164 NAC CUC1, CUC2, NAC1

miR167 ARF ARF6, ARF8

miR169 HAP At1g17590, At1g72830, At1g54160

miR171 SCL SCL6-III, SCL6-IV

miR172 AP2 AP2, TOE1, TOE2, TOE3

miR393 bZIP At1g27340

miR396 GRL GRL1, GRL2, GRL3, GRL7, GRL8

miR444 MADS Os02g49840

2.6. Regulatory Role of HD-ZIP III in Combination with

miR165/166

Based on sequence prediction, biochemical studies and genetic analysis, HD-ZIP III

genes have been proven to be targets of miR165/166 (Kim et al. 2005, Williams et al.

2005). MiR165/166 regulate PHB/PHV/REV mainly through mRNA cleavage (Emery

et a. 2003) and also promote DNA methylation of the PHB and PHV loci that is


30

likely to lead to transcriptional silencing of these genes (Bao et al. 2004). HD-ZIP III

genes and its regulator, miR165⁄166 genes have been found in gymnosperms, fern,

lycopod, moss, liverwort and hornwort, because they make an important contribution

to maintain the SAM and determine adaxial abaxial polarity in plant tissues. The

miR165 ⁄ 166 binding sites have been conserved in all HD-ZIP III genes of land plants

over hundreds of millions of years (Floyd and Bowman 2004). In contrast the HD-ZIP

III gene is also present in members of the Charales, a sister group of land plants and

five base changes are identified on the target site of miR165⁄166 in the HD-ZIP III

gene duplicated from Charales (Floyd et al. 2006; Sakaguchi and Watanabe 2012).

Thus, miRNA regulation appears to be absent from members of the Charales. Thus, it

was speculated that loss of complementarity on the HD-ZIP III coding sequences

hindered binding to miR165 ⁄ 166, and escape from the negative regulation triggered

by this binding. An early clue to such a regulatory network was given by Mallory et

al. (2004b) who introduced a silent base change in the miR165⁄166 complementary

site in PHB mRNA that caused adaxialization of leaves, like phb-d mutants, and

revealed that the 3’region of the miRNA complementary site plays a critical role in

the recognition of PHB mRNA by miR165 ⁄ 166.

In Arabidopsis, maize and tobacco, nucleotide changes in the START domain of HD-

ZIP III genes result in dominant mutations, due to a loss of miR165 or miR166

mediated regulation (McConnell et al. 2001; Emery et al. 3003; Zhong et al. 2004;

McHale and Koning, 2004; Juarez et al. 2004). It has been observed that

overexpression of miR166 by activation tagging leading to decreased mRNA levels of

ATHB-9/PHV, ATHB-14/PHB and ATHB-15 concomitantly causes a phenotype

reminiscent of the phv-phb-cna triple mutant (Kim et al. 2005, Williams et al. 2005).

Zhou et al. (2007) demonstrated that overexpression of miR165 cause a drastic

reduction in the mRNA levels of all five HD-ZIP III genes that results in loss-of-

function phenotypes including loss of SAM, altered organ polarity, defective vascular

development and impaired interfascicular fiber differentiation. Recent global gene

expression analysis revealed a link between miR165 overexpression and alteration in

the expression of genes involved in auxin signalling (Zhou et al. 2007). It has been

shown that a point mutation in the miR165 target sequence in IFL1/REV causes an


31

inhibition in the transcript cleavage and a high level accumulation of its full length

transcript (Zhong and Ye 2004) which leads to dominant phenotypes, including

formation of amphivasal vascular bundles and altered organ polarity (Zhong and Ye

1999; Emery et al. 2003; Zhong and Ye 2004). Likewise, miR165/166 is mobile in the

root and its mobility over a short distance is critical for dosage dependent regulation

of HD-ZIP III transcription factors in xylem patterning (Carlsbecker et al. 2010).

Studies have made it evident that HD-ZIP III and miR165/166 are also crucial factors

in flower development (Jung and Park 2007; Ji et al. 2011). Reduction in HD-ZIP III

expression by over-expression of miR165/166 and mis-expression of it by rendering

them resistant to miR165/166 leads to prolonged floral stem cell activity. This

indicates that the expression of HD-ZIP III genes needs to be precisely controlled to

achieve floral stem cell termination (Ji et al. 2011). Taken together these results

suggest that miR165 plays important roles in concert with miR166 in the regulation of

HD-ZIP III genes which have big impact in proper growth and development of plants.

The present survey of literature shows that there is little information available on

molecular analysis of developmental regulator in Brassica species. We therefore have

attempted to employ comparative genomics between Arabidopsis thaliana and

Brassica to analyse genomic re-arrangements in orthologous segments and to identify

transcription factor and miRNA genes. Subsequently, we have carried out detailed

sequence analysis and functional characterization through expression analysis and by

generating miRNA over-expressing transgenic plants in B. juncea genetic background

with an objective of understanding the role of this regulatory element in Brassica

development.

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2.1. Comparative Genomicsshodhganga.inflibnet.ac.in/bitstream/10603/12273/5/08_chapter 2.pdf · 2.1. Comparative Genomics In the post-genomic era, comparative analyses between genomes