M.Sc. I (Microbiology) MB-602:Molecular Biology Credit I RNA - - PowerPoint PPT Presentation

M.Sc. I (Microbiology) MB-602:Molecular Biology Credit –I

RNA Processing & Molecular Techniques

• Dr. Ashok Bankar

Assistant Professor, Department of Microbiology, MES-Abasaheb Garware College Affiliated to Savitribai Phule Pune University, Pune, Maharashtra, India. Email : ashokbankar@gmail.com ; ashokbankar@rediffmail.com

RNA Processing : Eukaryotic

When an RNA transcript is first made in a eukaryotic cell, it is considered a pre- mRNA and must be processed into a messenger RNA (mRNA). In humans and other eukaryotes, a freshly made RNA transcript is not quite ready to go. Instead, it's called a pre-mRNA and has to go through some processing steps to become a mature messenger RNA (mRNA) that can be translated into a protein. A 5' cap is added to the beginning of the RNA transcript, and a 3' poly-A tail is added to the end. In n splicing, some sections of the RNA transcript (introns) are removed, and the remaining sections (exons) are stuck back together. Some genes can be alternatively spliced, leading to the production of different mature mRNA molecules from the same initial transcript.

In bacteria, RNA transcripts are ready to act as messenger RNAs and get translated into proteins right away. In eukaryotes, things are a little more complex, though in an pretty interesting way. The molecule that's directly made by transcription in one

• f your (eukaryotic) cells is called a pre-mRNA, reflecting that it needs to go

through a few more steps to become an actual messenger RNA (mRNA). 1. Addition of a 5' cap to the beginning of the RNA 2. Addition of a poly-A tail (tail of A nucleotides) to the end of the RNA 3. Chopping out of introns, or "junk" sequences, and pasting together of the remaining, good sequences (exons) Once it's completed these steps, the RNA is a mature mRNA. It can travel out of the nucleus and be used to make a protein.

5' cap and poly-A tail :

Both ends of a pre-mRNA are modified by the addition of chemical groups. The group at the beginning (5' end) is called a cap, while the group at the end (3' end) is called a tail. Both the cap and the tail protect the transcript and help it get exported from the nucleus and translated on the ribosomes (protein-making "machines") found in the cytosol. The 5’ cap is added to the first nucleotide in the transcript during transcription. The cap is a modified guanine (G) nucleotide, and it protects the transcript from being broken down. It also helps the ribosome attach to the mRNA and start reading it to make a protein.

How is the poly-A tail added?

The 3' end of the RNA forms in kind of a bizarre way. When a sequence called a polyadenylation signal shows up in an RNA molecule during transcription, an enzyme chops the RNA in two at that site. Another enzyme adds about 100 - 200 adenine (A) nucleotides to the cut end, forming a poly-A tail. The tail makes the transcript more stable and helps it get exported from the nucleus to the cytosol.

RNA splicing : The third big RNA processing event that happens in your cells

is RNA splicing. In RNA splicing, specific parts

• f

the pre-mRNA, called introns are recognized and removed by a protein-and-RNA complex called the spliceosome. Introns can be viewed as "junk" sequences that must be cut out so the "good parts version" of the RNA molecule can be assembled. What are the "good parts"? The pieces of the RNA that are not chopped out are called exons. The exons are pasted together by the spliceosome to make the final, mature mRNA that is shipped out of the nucleus.

A key point here is that it's only the exons of a gene that encode a protein. Not only do the introns not carry information to build a protein, they actually have to be removed in order for the mRNA to encode a protein with the right sequence. If the spliceosome fails to remove an intron, an mRNA with extra "junk" in it will be made, and a wrong protein will get produced during translation.

Alternative splicing :

Why splice? We don't know for sure why splicing exists, and in some ways, it seems like a wasteful system. However, splicing does allow for a process called alternative splicing, in which more than one mRNA can be made from the same gene. Through alternative splicing, we (and other eukaryotes) can sneakily encode more different proteins than we have genes in our DNA. In alternative splicing, one pre-mRNA may be spliced in either of two (or sometimes many more than two!) different ways. For example, in the diagram below, the same pre-mRNA can be spliced in three different ways, depending on which exons are kept. This results in three different mature mRNAs, each of which translates into a protein with a different structure.

Processing of Eukaryotic Pre-mRNA :

Three major events occur during the process: 1) 5 capping 3) 3 cleavage/polyadenylation 3) RNA splicing 2) Processing occurs in The nucleus as the nascent mRNA precursor is being transcribed and the functional mRNA produced is transported to the cytoplasm. After nascent RNA molecules produced by RNA polymerase II reach a length of 25–30 nucleotides, 7-methylguanosine and the other components of the 5 cap found on eukaryotic mRNAs are added to their 5 end. This initial step in RNA processing is catalyzed by a dimeric capping enzyme, which associates with the phosphorylated carboxyl terminal domain (CTD) of RNA polymerase II. The CTD becomes phosphorylated during transcription initiation. Because the capping enzyme does not associate with polymerase I or III, which do not contain a CTD, capping is specific for transcripts produced by RNA polymerase II.

One subunit of the capping enzyme removes the γ phosphate from the 5 end of the nascent RNA emerging from the surface of an RNA polymerase II. Another domain of this subunit transfers the GMP moiety from GTP to the 5- diphosphate of the nascent transcript, creating the unusual guanosine 5’-5’- triphosphate structure. In the final steps, separate enzymes transfer methyl groups from S- adenosylmethionine to the N7 position of the guanine and the 2’ oxygens of riboses at the 5 end of the nascent RNA. From the time nascent transcripts first emerge from RNA polymerase II until mature mRNAs are transported into the cytoplasm, the RNA molecules are associated with an abundant set of nuclear proteins. Proteins are the major protein components of heterogeneous ribonucleoprotein particles (hnRNPs), which contain heterogeneous nuclear RNA (hnRNA), a collective term referring to pre-mRNA and other nuclear RNAs of various sizes.

The RNA recognition motif (RRM), also called the RNP motif and the RNA binding domain (RBD), is the most common RNA-binding domain in hnRNP proteins. This ≈80-residue domain, which occurs in many other RNA-binding proteins, contains two highly conserved sequences (RNP1 and RNP2) that allow the motif to be recognized in newly sequenced genes. The conserved RNP1 and RNP2 sequences lie side by side on the two central strands, and their side chains make multiple contacts with a single-stranded region

• f RNA.

The central RNP1 and RNP2 strands forming a positively charged surface that interacts with the negatively charged RNA phosphates.

Functions of hnRNP Proteins

The association of pre mRNAs with hnRNP proteins prevents formation of short secondary structures dependent on base-pairing of complementary regions, thereby making the pre-mRNAs accessible for interaction with other RNA molecules or proteins. Pre mRNAs associated with hnRNP proteins present a more uniform substrate for further processing steps than would free, unbound pre-mRNAs. Some hnRNP proteins may interact with the RNA sequences that specify RNA splicing or cleavage/polyadenylation and contribute to the structure recognized by RNA-processing factors. Some hnRNP proteins remain localized in the nucleus, whereas others cycle in and out of the cytoplasm, suggesting that they function in the transport of mRNA.

Splicing Occurs at Short, Conserved Sequences in Pre-mRNAs via Two Transesterification Reactions

During formation of a mature, functional mRNA, the introns are removed and exons are spliced together. For short transcription units, RNA splicing usually follows cleavage and polyadenylation of the 3’ end of the primary transcript. The location of splice sites—that is, exon-intron junctions—in a pre-mRNA can be determined by comparing the sequence of genomic DNA with that of the cDNA prepared from the corresponding mRNA Sequences that are present in the genomic DNA but absent from the cDNA represent introns and indicate the positions of splice sites. Such analysis of a large number of different mRNAs revealed moderately conserved, short consensus sequences at the splice sites flanking introns in eukaryotic pre-mRNAs ; in higher organisms, a pyrimidine-rich region just upstream of the 3’ splice site also is common. Studies with deletion mutants have shown that much of the center portion of introns can be removed without affecting splicing; generally only 30–40 nucleotides at each end of an intron are necessary for splicing to occur at normal rates.

Analysis of the intermediates formed during splicing of pre-mRNAs in vitro led to the discovery that splicing

• f

exons proceeds via two sequential transesterification reactions. Introns are removed as a lariat-like structure in which the 5’G of the intron is joined in an unusual 2,5-phosphodiester bond to an adenosine near the 3’ end of the intron. This A residue is called the branch point because it forms an RNA branch in the lariat structure. In each transesterification reaction, one phosphoester bond is exchanged for another. Since the number of phosphoester bonds in the molecule is not changed in either reaction, no energy is consumed. The net result of these two reactions is that two exons are ligated and the intervening intron is released as a branched lariat structure.

Five U-rich small nuclear RNAs (snRNAs), designated U1, U2, U4, U5, and U6, participate in pre-mRNA splicing. Ranging in length from 107 to 210 nucleotides, these snRNAs are associated with 6 to 10 proteins in small nuclear ribonucleoprotein particles (snRNPs) in the nucleus

• f eukaryotic cells.

Definitive evidence for the role of U1 snRNA in splicing came from experiments which indicated that base pairing between the 5 splice site of a pre-mRNA and the 5 region of U1 snRNA is required for RNA splicing

Involvement of U2 snRNA in splicing initially was suspected when it was found to have an internal sequence that is largely complementary to the consensus sequence flanking the branch point in pre-mRNAs. Compensating mutation experiments, similar to those conducted with U1 snRNA and 5 splice sites, demonstrated that base pairing between U2 snRNA and the branch-point sequence in pre-mRNA also is critical to splicing. the general structures of the U1 and U2 snRNAs and how they base-pair with pre- mRNA during splicing. Significantly, the branch-point A itself, which is not base-paired to U2 snRNA, “bulges out,” allowing its 2 hydroxyl to participate in the first transesterification reaction of RNA splicing

Spliceosomes, Assembled from snRNPs and a Pre-mRNA, Carry Out Splicing

According to the current model of pre-mRNA splicing, the five splicing snRNPs are thought to assemble on the premRNA, forming a large ribonucleoprotein complex called a spliceosome. Assembly of a spliceosome begins with the base pairing of the snRNAs of the U1 and U2 snRNPs to the pre-mRNA. Extensive base pairing between the snRNAs in the U4 and U6 snRNPs forms a complex that associates with U5 snRNP. The U4/U6/U5 complex then associates with the previously formed U1/U2/pre- mRNA complex to yield a spliceosome.

After formation of the spliceosome, extensive rearrangements in the pairing of snRNAs and the pre-mRNA lead to release of the U1 and U4 snRNPs. The catalytically active rearranged spliceosome then mediates the first transesterification reaction that forms the 2,5-phosphodiester bond between the 2 hydroxyl on the branch point A and the phosphate at the 5 end of the intron.

Following another rearrangement of the snRNPs, the second transesterification reaction ligates the two exons in a standard 3,5-phosphodiester bond, releasing the intron as a lariat structure associated with the snRNPs. This final intron-snRNP complex rapidly dissociates, and the individual snRNPs released can participate in a new cycle of splicing. The excised intron is then rapidly degraded by a debranching enzyme and other nuclear RNases.

Self-Splicing Group II Introns

Under certain non physiological in vitro conditions, pure preparations of some RNA transcripts slowly splice out introns in the absence of any protein. This

• bservation led to recognition that some introns are self-splicing.

Two types of self-splicing introns have been discovered: group I introns, present in nuclear rRNA genes of protozoans, and group II introns, present in protein- coding genes and some rRNA and tRNA genes in mitochondria and chloroplasts

• f plants and fungi.

all group II introns fold into a conserved, complex secondary structure containing numerous stem-loops. Self-splicing by a group II intron occurs via two transesterification reactions, involving intermediates and products analogous to those found in nuclear pre- mRNA splicing. The mechanistic similarities between group II intron self-splicing and spliceosomal splicing led to the hypothesis that snRNAs function analogously to the stem-loops in the secondary structure of group II introns.

According to this hypothesis, snRNAs interact with 5 and 3 splice sites of pre mRNAs and with each other to produce a three-dimensional RNA structure functionally analogous to that of group II self-splicing introns.

An extension of this hypothesis is that introns in ancient pre-mRNAs evolved from group II self-splicing introns through the progressive loss of internal RNA structures, which concurrently evolved into trans-acting snRNAs that perform the same functions. Support for this type of evolutionary model comes from experiments with group II intron mutants in which domain V and part of domain I are deleted. RNA transcripts containing such mutant introns are defective in self-splicing, but when RNA molecules equivalent to the deleted regions are added to the in vitro reaction, selfsplicing occurs. This finding demonstrates that these domains in group II introns can be trans- acting, like snRNAs. The similarity in the mechanisms of group II intron self splicing and spliceosomal splicing of pre-mRNAs also suggests that the splicing reaction is catalyzed by the snRNA, not the protein, components of spliceosomes.

Although group II introns can self-splice in vitro at elevated temperatures and Mg2+ concentrations, under in vivo conditions proteins called maturases, which bind to group II intron RNA, are required for rapid splicing. Maturases are thought to stabilize the precise three-dimensional interactions of the intron RNA required to catalyze the two splicing transesterification reactions. By analogy, snRNP proteins in spliceosomes are thought to stabilize the precise geometry of snRNAs and intron nucleotides required to catalyze pre-mRNA splicing.

Group I Intron

Processing of rRNA and tRNA

Approximately 80 percent of the total RNA in rapidly growing mammalian cells (e.g., cultured HeLa cells) is rRNA, and 15 percent is tRNA; protein-coding mRNA thus constitutes only a small portion of the total RNA. The primary transcripts produced from most rRNA genes and from tRNA genes, like pre-mRNAs, are extensively processed to yield the mature, functional forms

• f these RNAs.

Pre-rRNA Genes Are Similar in All Eukaryotes

The 28S and 5.8S rRNAs associated with the large (60S) ribosomal subunit and the 18S rRNA associated with the small (40S) ribosomal subunit in higher eukaryotes are encoded by a single type of pre-rRNA transcription unit. Transcription by RNA polymerase I yields a 45S (13.7-kb) primary transcript (pre-rRNA), which is processed into the mature 28S, 18S, and 5.8S rRNAs found in cytoplasmic ribosomes.

Cloning and sequencing of the DNA encoding pre-rRNA from many species showed that this DNA shares several properties in all eukaryotes. First, the pre-rRNA genes are arranged in long tandem arrays separated by non transcribed spacer regions ranging in length from ≈2 kb in frogs to ≈30 kb in humans. Second, the genomic regions corresponding to the three finished rRNAs are always arranged in the same 5′ → 3′ order: 18S, 5.8S, and 28S

Third, in all eukaryotic cells (and even in bacteria), the pre-rRNA gene, as well as the corresponding primary transcript, is considerably longer than the sum of the three finished rRNA molecules. For example, in human cells only about half of the 45S pre-rRNA primary transcript appears in the final rRNA products, whose combined length is about 7.2

• kb. The other half, called transcribed spacer RNA, is removed during processing

and is rapidly degraded. Discovery of pre-rRNA processing was the first indication that mature cytoplasmic RNAs are derived from larger precursor RNAs synthesized in the nucleus. Both the synthesis and processing of pre-mRNA occurs in the nucleolus. When pre-rRNA genes initially were identified in the nucleolus by in situ hybridization, it was not known whether any other DNA was required to form the nucleolus. Subsequent experiments with transgenic Drosophila strains demonstrated that a single complete pre-rRNA transcription unit induces formation of a small nucleolus.

Thus a single pre-rRNA gene is sufficient to be a nucleolar organizer, and all the

• ther components of the ribosome diffuse to the newly formed pre-rRNA.

The structure of the induced nucleolus appears, at least by light microscopy, to be the same as, except smaller than, a normal Drosophila nucleolus containing 200 or so pre-rRNA genes.

Small Nucleolar RNAs (snoRNAs) Assist in Processing rRNAs and Assembling Ribosome Subunits

Following their synthesis in the nucleolus, nascent pre-rRNA transcripts are immediately bound by proteins, forming pre-ribonucleoprotein particles, or pre- rRNPs. Several ribonucleoprotein particles of different sizes have been extracted from mammalian nucleoli. The largest of these (80S) contains an intact 45S pre-rRNA molecule, which is cut in a series of cleavage and exonucleolytic steps that ultimately yield the mature rRNAs found in ribosomes

During processing, pre-rRNA also is extensively modified, mostly by methylation

• f the 2′-hydroxyl group of specific riboses and conversion of specific uridine

residues to pseudouridine. Some of the proteins in the pre-rRNPs found in nucleoli remain associated with the mature ribosomal subunits, whereas others are restricted to the nucleolus and assist in assembly of the subunits.

The positions of cleavage sites in pre-rRNA and the specific sites of 2′-O- methylation and pseudouridine formation are determined by approximately 150 different small nucleolus-restricted RNA species, called small nucleolar RNAs (snoRNAs), which hybridize transiently to pre-rRNA molecules. Like snRNAs, snoRNAs associate with proteins, forming snoRNPs. One large class of snoRNAs, involved in 2′-O-methylation, contain common sequences bound by the nucleolus-restricted protein fibrillarin. A conserved sequence in these snoRNAs, which is invariably positioned close to methylation sites in the pre-rRNA, is thought to bind a methyltransferase enzyme that modifies the ribose moiety. Another snoRNP, called RNase MRP, catalyzes one of the cleavages by which transcribed spacer sequences are removed from pre-rRNA. The associated snoRNA is homologous to the RNA of RNase P involved in tRNA

• processing. Based on this homology, the cleavage reaction is thought to be

catalyzed by the MRP snoRNA. There is strong evidence that RNase P performs

• ne of the pre-rRNA cleavages as well.

Some snoRNAs are expressed from their own promoters by RNA polymerase II or

• III. Remarkably, however, the large majority of snoRNAs are spliced-out introns
• f genes encoding functional mRNAs.

Unlike pre-rRNA genes, 5S-rRNA genes are transcribed by RNA polymerase III in the nucleoplasm outside of the nucleolus. Without further processing, 5S RNA diffuses to the nucleolus, where it assembles with the 28S and 5.8S rRNAs and proteins into large ribosomal subunits. When assembly of ribosomal subunits in the nucleolus is complete, they are transported through nuclear pore complexes to the cytoplasm, where they appear first as free subunits.

The DNA in the protozoan Tetrahymena thermophila contains an intervening intron in the region that encodes the large pre-rRNA molecule. Careful searches failed to uncover even one pre-rRNA gene without the extra sequence, indicating that splicing is required to produce mature rRNA in these organisms. Subsequent studies showing that the pre-rRNA was spliced at the correct sites when incubated by itself, without assistance from any protein, provided the first indication that RNA can function as a catalyst, like enzymes. Following the discovery of self-splicing in Tetrahymena pre-rRNA, a whole raft of self-splicing sequences were found in pre-rRNAs from other single-celled

• rganisms, in mitochondrial and chloroplast pre-rRNAs.

The self-splicing sequences in all these precursors, referred to as group I introns, use guanosine as a cofactor and can fold by internal base pairing to juxtapose closely the two exons that must be joined. Clearly, in the self-splicing introns, RNA functions as a ribozyme, an RNA sequence with catalytic ability.

The group I intron within the pre-rRNA of Tetrahymena and certain other organisms is unrelated to the transcribed spacer sequences that separate the 18S, 5.8S, and 28S regions in the majority of organisms. In particular, the self-splicing mechanism that removes group I introns differs from the cleavage mechanism by which spacer sequences are removed during processing of pre-rRNA Mutational and biochemical experiments are under way to determine which residues are critical in catalyzing the two transesterification reactions leading to splicing.

All Pre-tRNAs Undergo Cleavage and Base Modification

Mature cytosolic tRNAs, which average 75 – 80 nucleotides in length, are produced from larger precursors (pre-tRNAs) synthesized by RNA polymerase III in the nucleoplasm. Mature tRNAs also contain numerous modified bases that are not present in tRNA primary transcripts. Cleavage and base modification occur during processing of all pre-tRNAs. Some pre-tRNAs contain one or more introns that are spliced out during processing. A 5′ sequence of variable length that is absent from mature tRNAs is present in all pre-tRNAs .

These extra 5′ nucleotides are removed by the ribonuclease P (RNase P), a ribonucleoprotein endonuclease. The RNase P polypeptide increases the rate of cleavage by M1 RNA, allowing it to proceed at physiological Mg2+ concentrations.

A 14-nucleotide intron (blue) in the anticodon loop is removed by splicing. A 16- nucleotide sequence (green) at the 5′ end is cleaved by RNase P. U residues at the 3′ end are replaced by the CCA sequence (red) found in all mature tRNAs. Numerous bases in the stem-loops are converted to characteristic modified bases (yellow). Not all pre-tRNAs contain introns that are spliced out during processing, but they all undergo the other types of changes shown here. D = dihydrouridine; Ψ = pseudouridine.0

About 10 percent of the bases in pre-tRNAs are modified enzymatically during

• processing. Three types of base modifications occur : 1) replacement of U residues

at the 3′ end of pre-tRNA with a CCA sequence, which is found at the 3′ end of all tRNAs 2) addition of methyl and isopentenyl groups to the heterocyclic ring of purine bases and methylation of the 2′-OH group in the ribose of any residue; and 3) conversion

• f

specific uridines to dihydrouridine, pseudouridine,

• r

ribothymidine residues. Processing of pre-tRNA, like mRNA processing, occurs in the nucleoplasm. The mature tRNAs then are transported to the cytoplasm through nuclear pore complexes. Interestingly, U6 snRNA, another RNA synthesized by RNA polymerase III, is a component of the spliceosome and remains in the nucleus, whereas tRNAs are efficiently transported to the cytoplasm. Most likely, mature tRNAs in the nucleus, like mature mRNAs and rRNAs, are bound by specific proteins that facilitate their transport through nuclear pores. Once in the cytoplasm, tRNAs are passed between aminoacyl-tRNA synthetases, elongation factors, and ribosomes during protein synthesis. Thus tRNAs generally are associated with proteins and spend little time free in the cell, as is also the case for mRNAs and rRNAs.

Splicing of Pre-tRNAs Differs from Other Splicing Mechanisms

Comparison of the sequences of tRNA genes with the sequences of the corresponding cytosolic tRNAs has shown that some eukaryotic nuclear tRNA genes contain introns. For example, the pre-tRNA expressed from the yeast tyrosine tRNA (tRNA

Tyr) gene contains a 14-base intron that is not present in

mature tRNATyr Some archaeal tRNA genes also contain introns. The introns in nuclear pre- tRNAs are shorter than those in pre-mRNAs, and they do not contain the splice- site consensus sequences found in pre-mRNAs. Pre-tRNA introns also are clearly distinct from the much longer self-splicing group I and group II introns found in chloroplast and mitochondrial pre- rRNAs. The mechanism of pre-tRNA splicing, outlined in Figure 11-53, differs in several ways from the mechanisms utilized by self-splicing introns and spliceosomes.

For instance, during pre-tRNA splicing the intron is excised in one step rather than two; GTP and ATP are required; a 2′, 3′-cyclic monophosphate forms on the cleaved end of the 5′ exon; and the process is catalyzed by proteins (enzymes) rather than RNA. Certain mutations in pre-tRNA that change its secondary structure prevent the splicing reaction, indicating that pre-tRNA molecules must be folded into a particular secondary structure for intron excision to occur. Since introns always are found in the anticodon loop of pre-tRNAs, pre-tRNAs most likely are folded similarly to mature tRNAs, thereby bringing the two intron-exon junctions into proximity. When intron-containing yeast tRNA genes are microinjected into Xenopus oocyte nuclei, correctly processed tRNAs are produced. This finding indicates that enzymatic systems for cleaving, modifying, and splicing pre-tRNAs have been conserved over a wide evolutionary range.

First, the pre-tRNA is cleaved at two places, on each side of the intron, thereby excising the intron. The cleavage mechanism generates a 2′,3′-cyclic phosphomonoester at the 3′ end of the 5′ exon. The multistep reaction joining the two exons requires two nucleoside triphosphates: a GTP, which contributes the phosphate group (yellow) for the 3′ → 5′ linkage in the finished tRNA molecule; and an ATP, which forms an activated ligase-AMP intermediate. The 2′- phosphate on the 5′ exon is removed in the final step

RNA editing can be broadly defined as any site-specific alteration in an RNA sequence that could have been copied from the template, excluding changes due to processes such as RNA splicing and polyadenylation. Changes in gene expression attributed to editing have been described in organisms from unicellular protozoa to man, and can affect the mRNAs, tRNAs, and rRNAs present in all cellular compartments. These sequence revisions, which include both the insertion and deletion of nucleotides, and the conversion of one base to another, involve a wide range of largely unrelated mechanisms.

RNA EDITING

Editing of adenosine (A) to inosine (I) in double-stranded RNA, catalyzed by adenosine deaminases acting on RNA (ADARs), is one dynamic modification that in a combinatorial manner can give rise to a very diverse transcriptome. Since the cell interprets inosine as guanosine (G), editing can result in non- synonymous codon changes in transcripts as well as yield alternative splicing, but also affect targeting and disrupt maturation of microRNA. ADAR editing is essential for survival in mammals but its dysregulation can lead to cancer. ADAR1 is for instance over expressed in, e.g., lung cancer, liver cancer, esophageal cancer and chronic myoelogenous leukemia, which with few exceptions promotes cancer progression. In contrast, ADAR2 is lowly expressed in e.g. glioblastoma, where the lower levels of ADAR2 editing leads to malignant phenotypes. Altogether, RNA editing by the ADAR enzymes is a powerful regulatory mechanism during tumorigenesis.

ADARs

MECHANISMS OF INSERTION/DELETION EDITING

The internal insertion of nucleotides has been observed in mitochondrial RNAs from kinetoplastid and amoebid protozoa, myxomycetes, chytriomycete fungi, and nematodes. Indeed, insertional editing can

• ccur

either post-transcriptionally

• r

cotranscriptionally. 1. Posttranscriptional Nucleotide Insertion/Deletion The term RNA editing was first coined by Benne and colleagues to describe the insertion of uridines into the cytochrome oxidase subunit II mRNA in kinetoplasts of Trypanosoma brucei and Crithidia fasciculata. The global nature of such frame shifting events within the kinetoplastid protozoa was soon established by the identification of additional examples in these species and in Leishmania tarentolae and later extended to more distant relatives.

Deletion of uridine residues was also observed, albeit at lower frequency, within each of these species. The extent of editing was shown to vary from the insertion of a few nucleotides to extensive insertion/deletion of uridine residues (pan-editing) in which over 50% of the final mRNA product is the result of RNA editing. The notion of “editing” sequence at the RNA level was originally met with skepticism, based largely on the absence of any obvious template that could be Used to direct uridine insertion/deletion. The identification of guide RNAs (gRNAs) as the likely source of the missing information quickly led to the formulation of two general models by which accurate insertion and deletion of nucleotides could be achieved. These models, cleavage-ligation and transesterification, made different predictions as to the expected reaction intermediates and stimulated a flurry of Experimentation and debate. Development of in vitro systems capable of carrying

• ut uridine deletion (177, 178) and insertion (36, 49, 94) permitted direct testing of

particular features of individual models, resulting in the eventual acceptance of the cleavage-ligation pathway as the mechanism of kRNA editing (4, 73, 195).

Transport of messenger RNA (mRNA) from the nucleus to the cytoplasm is an essential step of eukaryotic gene expression. In the cell nucleus, a precursor mRNA undergoes a series of processing steps, including capping at the 5' ends, splicing and cleavage/polyadenylation at the 3' ends. During this process, the mRNA associates with a wide variety of proteins, forming a messenger ribonucleoprotein (mRNP) particle. Association with factors involved in nuclear export also occurs during transcription and processing, and thus nuclear export is fully integrated into mRNA maturation. The coupling between mRNA maturation and nuclear export is an important mechanism for providing only fully functional and competent mRNA to the cytoplasmic translational machinery, thereby ensuring accuracy and swiftness of gene expression.

Nuclear export of mRNA

The molecular mechanism of nuclear mRNA export mediated by the principal transport factors, including Tap-p15 and the TREX complex. Nuclear pore complexes (NPCs), which perforate the NE, are the main gateways through which RNAs and proteins are delivered to their proper destinations. The NPC is composed of approximately 30 distinct proteins that are collectively known as nucleoporins. A subset of nucleoporins that line the central transport channel contains phenylalanine-glycine (FG)-repeat sequences, which emanate to the inside of the channel and form a dense hydrophobic meshwork that functions as a barrier limiting the improper exchange of soluble macromolecules between the nucleus and the cytoplasm. Thus, nucleo-cytoplasmic transport of RNAs and proteins requires specific transport receptors to break this barrier. The importin/karyopherin-β family of proteins comprise the prototypical transport receptor family that mediates nucleo-cytoplasmic movement of most proteins and small non-coding RNAs, such as tRNA, uridine-rich small nuclear RNA (UsnRNA), and miRNA.

Nuclear export of mRNAs is a unique process that does not directly rely on the functions of the importin/karyopherin-β transport receptor family and Ran. Instead, it requires the evolutionarily conserved heterodimeric transport receptors Tap-p15 (also called Nxf1-Nxt1) in metazoans and Mex67-Mtr2 in yeast. Both Tap-p15 and Mex67-Mtr2 are RNA binding proteins, but they bind nonspecifically to RNA in vitro and are not able to distinguish different RNAs on their own. To circumvent this problem, a series of mRNA-binding proteins participate in this

• process. The conserved transcription-export (TREX) complex, which consists of

the THO subcomplex and Uap56 and Aly/REF plays an important role in selection

• f mRNAs by Tap-p15 and Mex67-Mtr2.

The RNA-binding components of the TREX complex, including yeast Yra1 and mammalian Aly/REF, directly interact with the export receptor heterodimers, thereby functioning as adaptors.

In addition, in yeast, the serine-arginine rich (SR) proteins Npl3, Gbp2 and Hrb1are associated with the TREX complex and the mRNA binding protein Nab2 also interact with Mex67-Mtr2 and probably function as adaptors. In mammalian cells, the SR proteins 9G8 and SRp20, as well as numerous mRNA-binding proteins, have been proposed to play a similar role. Recruitment of adaptor proteins to mRNPs is coupled with transcription and processing, causing mRNPs to be licensed to the mRNA-specific export pathway upon the completion of nuclear processing. Thus, transcription by RNA polymerase II (RNAPII) is a key determinant allocating mRNA to the appropriate export pathway.

During transcription, protein factors required for capping, splicing and cleavage/polyadenylation are recruited to the nascent transcript, forming an mRNP. The 5' end of the mRNA is capped early in this process via an interaction between the capping enzyme and RNA polymerase II (RNAPII). Factors involved in splicing and cleavage/polyadenylation are also co- transcriptionally loaded onto the pre-mRNA. The TREX complex and a subset of the SR proteins, which are engaged in nuclear export, are recruited to the nascent mRNA via interactions with the transcription and processing factors. The nuclear export receptor Tap-p15 (Mex67-Mtr2 in yeast) in turn gains access to the mRNA via interactions with these factors as adaptors. The nuclear export receptor heterodimer facilitates the translocation of mRNPs through its interaction with FG-repeat containing nucleoporins.

During the process of the nuclear mRNA biogenesis, the structure and the composition of the mRNP change drastically and these alterations in the physicochemical properties also help the mRNP translocate through the NPC. The mRNA export factors are then dissociated from the mRNP by factors associated with the NPC to prevent the return of the mRNP to the nucleus. The exported mRNA then directs protein translation in the cytoplasm.

Regulatory RNA And Non Coding RNA (nc RNA)

Though 80% of the human genome is transcribed into RNA, majority of RNA lacks protein coding potential and referred as “non-coding RNA” (ncRNA). The mammalian transcriptome is much more complex and their transcription is regulated by developmental stages. The continuing discovery of new classes of regulatory ncRNAs suggests that RNA has continued to evolve along with proteins and DNA. The ncRNAs are divided into two major groups based on an arbitrary threshold of 200 nucleotides (nt) namely 1) short ncRNAs (sncRNA) and 2) long ncRNAs (lncRNAs) . The sncRNAs include functional RNAs such as t-RNAs, r-RNAs and snRNAs which are involved in transcriptional and translational regulation. The short ncRNAs also include different regulatory RNAs such as microRNAs (miRNAs), small interfering RNAs (siRNAs) and P-element-induced wimpy testis (PIWI) interacting RNAs (piRNAs), all of which regulate gene expression.

miRNAs

The miRNAs are 20–30 nucleotides long and generated from sense and antisense DNA strands. MicroRNAs are found in the plant and animal branches of Eukaryota and are encoded by a bewildering array of genes. They induce mRNA degradation or translational repression, which in turn result in the alteration of gene expression. About 60% of translated protein coding genes are negatively regulated by miRNAs. Some transcripts are regulated by a single miRNA, while others are regulated by more than one miRNAs. In addition to the transcriptional gene regulation, miRNAs play important roles in pivotal biological processes such as cell proliferation, cell differentiation, development, and cell death.

The miRNA biogenesis and mechanism of action

The process of miRNA biogenesis is quite characteristic for the ncRNAs subclass. Based on cellular requirement, the primary miRNA transcript (pri-miRNA) is first transcribed from the DNA by RNA polymerase II and transcripts are capped and

• polyadenylated. They are characterized by one or many stem-loop hairpins which

encompass the functional mature miRNA in their stem. In animals, the first step occurs in nucleus, in which the pri-miRNA upon recognition by two nuclear enzymes, Drosha and DGCR8 is processed into dsRNA molecule containing one or more hairpins of approximately 70 nucleotides long, which are called as precursor miRNAs (pre-miRNAs). Then they are exported to the cytoplasm by the nuclear export protein exportin-5. In cytoplasm, the pre-miRNA is recognized and processed by the RNase III enzyme, Dicer which removes the hairpin loop resulting in 20–23 nt dsRNA (miRNA-miRNA*) molecule. When the complementarity between the miRNA bound to Argonaute protein 1 (Ago1) protein and the target m-RNA is high, miRNA tailing and 3′–5′ trimming

• ccurs.

The RNA induced silencing (RISC) complex then targets the mRNA transcript based on sequence complementarity between the miRNA sequence and nucleotides in the 3′ untranslated regions (3′ UTR) of the target mRNAs. The binding of the the RNA-induced silencing complex (RISC) to its target leads to direct Ago- mediated cleavage of the target and causes mRNA degradation if the homology between miRNA and its target mRNA is extensive. Initially, it has been showed that miRNAs mainly target the 3′ UTRs of mRNAs, but recently, it was found that miRNA target sites also been located in the 5′ UTRs and even in coding regions of some of the target mRNAs. For example, mir-148 targets

• n the coding regions of DNMT3B.

Non repressed mRNAs recruit initiation factors and ribosomal subunits and form circularized structures that enhance translation (top). When miRISCs bind to mRNAs, they can repress initiation at the cap recognition stage (upper left) or the 60S recruitment stage (lower left). Alternatively, they can induce deadenylation of the mRNA and thereby inhibit circularization of the mRNA (bottom). They can also repress a postinitiation stage of translation by inducing ribosomes to drop off prematurely (lower right). Finally, they can promote mRNA degradation by inducing deadenylation followed by decapping.

siRNA

The canonical inducer of RNAi is long, linear, perfectly base paired dsRNA, introduced directly into the cytoplasm or taken up from the environment. These dsRNAs are processed by Dicer into the siRNAs that direct silencing. The siRNAs were originally observed during trans gene and virus-induced silencing in plants, consistent with a natural role in genome defense. In 2002 and 2003, centromeres, transposons, and other repetitive sequences were uncovered as another wellspring of siRNAs.

siRNA Biogenesis and Mechanism

In cytoplasm, the small RNA duplex molecules produced by the action of Dicer, creates a RNA duplexes with 2-nt overhangs at their 3′ ends and phosphate groups at their 5′ ends. Only one of the two strands of dsRNA acts as a guide strand and directs gene- silencing while, the other strand incorporates into the RNA-induced silencing complex (RISC) containing the Argonaute proteins (Ago2) and the GW182. The siRNAs are recognized by Argonaute protein 2 (Ago2). The pathways have been characterized in Drosophila and in humans. The siRISC assembly in Drosophila is nucleated by the R2D2/Dicer-2 heterodimer, which binds an siRNA duplex and then progresses by the addition of unknown factors to form the RISC-loading complex (RLC). The RLC then assembles into pre-RISC, with the siRNA still in duplex form. The pre-RISC formation is the first step that requires Ago2, the Drosophila Ago protein. The Ago2 then cleaves the passenger strand, leading to its ejection and the conversion of the entire assembly into the 80S holo-RISC.

RISC assembly in humans has also been characterized biochemically and appears to be a simpler process. Three proteins—Dicer, TRBP, and Ago-2— associate with each other even in the absence of the dsRNA trigger. This trimer, also referred to as the RISC-loading complex, is capable of binding dsRNA, dicing it into an siRNA, loading the siRNA into Ago-2, and discarding the passenger strand to generate functional RISC. Additional proteins associate with Ago complexes from human cells, but they do not appear to be essential for RISC loading or target cleavage. In many species, the siRNA populations that engage a target can be amplified by the action

• f

RNA-dependent RNA polymerase (RdRP) enzymes, strengthening and perpetuating the silencing response.

Different categories of siRNAs can depend upon different proteins for their function, indicating that they rely on different biogenesis and RISC assembly

• pathways. This is particularly true in plants, where viral siRNAs, transgene

siRNAs, and tasiRNAs have highly distinct cofactor requirements. During the canonical RNAi pathway, the siRNA guide strand directs RISC to perfectly complementary RNA targets, which are then degraded. The RNA degradation is induced by the PIWI domain of the Ago protein. This ‘‘slicer’’ activity is very precise : the phosphodiester linkage between the target nucleotides that are base paired to siRNA residues 10 and 11 is cleaved to generate products with 5’-monophosphate and 3’-hydroxyl termini. Once the initial cut is made, cellular exonucleases attack the fragments to complete the degradative process. The newly generated 3’ end of RISC cleavage products is also a substrate for

• ligouridylation,

which can promote exonucleolytic targeting. The target dissociates from the siRNA after cleavage, freeing RISC to cleave additional targets. In some cases, highly purified forms of RISC fail to cleave their targets with multiple turnover, suggesting that extrinsic factors promote product release, which is likely to be driven by ATP hydrolysis.

RNA i

RNA interference (RNAi) or Post-Transcriptional Gene Silencing (PTGS) is a conserved biological response to double-stranded RNA that mediates resistance to both endogenous parasitic and exogenous pathogenic nucleic acids, and regulates the expression of protein-coding genes. This natural mechanism for sequence-specific gene silencing promises to revolutionize experimental biology and may have important practical applications in functional genomics, therapeutic intervention, agriculture and other areas. Investigations on diverse organisms, labeled variously as PTGS in plants, RNAi in animals, quelling in fungi, and virus-induced gene silencing, have converged

• n a universal paradigm of gene regulation.

The critical common components of the paradigm are that (i) the inducer is the dsRNA (ii) the target RNA is degraded in a homology dependent fashion (iii) the degradative machinery requires a set of proteins which are similar in structure and function across most organisms.

In most of these processes, certain invariant features are observed, including the formation of small interfering RNA (siRNA) and the organism-specific systemic transmission of silencing from its site of initiation.

Mechanism of silencing

The RNAi-mediated gene silencing is executed by siRNAs. The process of silencing begins with the cleavage of long dsRNAs into 21–25 -nt fragments of siRNAs in cytoplasm . The process is catalyzed by Dicer enzyme. These siRNAs are inserted into multiprotein silencing complex, which is known as RNA-induced silencing complex (RISC). Subsequent unwinding of siRNA duplex, in turn, leads to active confirmation of RISC complex (RISC*). Next, target mRNA (mRNA to be degraded) is recognized by antisense RNA, which signals RISC complex for the endonucleolytic degradation of the homologous mRNA.

The target mRNA is cleaved in the centre of the region that is recognized by complimentary guide siRNA, which is 10–12 -nt away from the 5′ terminus of siRNA. The RNAi process is completed by the last step of siRNA molecule amplification. The siRNAs is derived from the priming on the target mRNA by RNA- dependent RNA polymerase (RdRp) enzyme by existing siRNAs. The second generation of siRNAs is effective in inducing a secondary RNA interference that is defined as transitive RNAi. The transitive RNAi causes a systemic genetic interference in plants and C. elegans. Interestingly, transitive and systemic RNAi is absent in Drosophila and mammals owing to the lack of RdRp in both organisms.

The Figure demonstrates that double-stranded RNA (dsRNA) can generate either from exogenous natural sources, such as a viral infection, exogenous artificial sources such as transfection, or natural synthesis. The dsRNA is then processed by a multimeric Dicer enzyme to generate siRNA that can be further amplified by RNA-dependent RNA polymerase (RdRp). The siRNA subsequently interacts with an array of proteins to form RNA-induced silencing complex (RISC) that is activated in an ATP-dependent manner. The activated RISC (RISC*) can then induce chromatin remodeling or TGS, or induce target RNA cleavage,

• r

cause miRNA-mediated translational inhibition.

PiRNAs (Piwi-interacting RNAs)

The piRNAs have emerged as an extremely complex population of small RNAs that are highly enriched in the germline tissues of the majority of metazoans. The Piwi/piRNA pathways are known to play roles in the fertility of diverse animal species, as evidenced by the fertility defects in mutants lacking Piwi. The piRNA function is the silencing of mobile elements. Mature piRNA sequences are surprisingly diverse between different organisms, even between closely related species. However, they share a number of characteristics other than their interaction with Piwi proteins. For example, in both D. melanogaster and vertebrates, piRNAs are between 26 and 30 nucleotides (nt) in length, have a ‘preference’ for a 5′ uracil, and posses a 3′-most sugar that is 2′-O-methylated. The piRNA biogenesis pathways in different organisms also appear to be diverse, and are distinct from those that produce miRNAs or siRNAs.

piRNA biogenesis in D. melanogaster

The D. melanogaster genome encodes three Piwi proteins 1) Piwi 2) Aubergine (Aub) and 3) Ago3. These all are required for male and female fertility. These Piwi proteins show distinct expression patterns : Piwi localises to nuclei in germ cells and the somatic follicle cells of the ovary. Aub is expressed in the cytoplasm of germ cells and localises partially to the nuage that plays a prominent role in piRNA function. The Ago3 is restricted to the cytoplasm of germ cells and is distinctly localised to the nuage. Different Piwi proteins bind to distinct sets of small RNAs : Piwi and Aub show a strong preference for sequences with a 5′ uridine (U) mapping antisense to transposons, whereas Ago3 piRNAs show no enrichment for 5′ U and are sense to transposons. Aub- and Ago3-associated piRNAs are generated by an amplification loop involving these two Piwi proteins. Aub-bound piRNAs recognise a complementary transposon transcript and induce endonucleolytic cleavage – slicing – of the target at position 10-11 of the piRNA.

Such slicing generates the 5′ end of a new sense piRNA with a 10 nt 5′ overlap with the initial antisense piRNA and an adenosine residue at position 10. After 3′ end processing and modification, this new piRNA is incorporated into Ago3 and goes on to generate Aub-bound piRNAs from piRNA cluster transcripts using the same mechanism. This cytoplasmic loop, which is also found in mice and zebrafish is referred to as the ‘ping-pong’ cycle, and links piRNA amplification to post-transcriptional target silencing.

Mechanism of piRNA-mediated transcriptional silencing

Mobile elements are the most prominent piRNA targets, and the cytoplasmic ping-pong cycle, in which a transposon target is recognised by a piRNA and sliced by the Piwi protein via its catalytic domain to give rise to a new piRNA with opposite orientation, is a particularly well-understood post-transcriptional mechanism for transposon repression. Many members of the Piwi protein family have a conserved catalytic domain and are therefore capable of target ‘slicing’. The identification of the ping-pong cycle in D. melanogaster and mice as an efficient means for both transposon transcript degradation and small RNA amplification clearly showed the requirement for a cytoplasmic component in piRNA silencing. Nevertheless, nuclear localisation of D. melanogaster Piwi and murine MIWI2 provided strong evidence for additional modes of silencing. A role for transcriptional silencing as a mechanism for piRNA-mediated silencing of transposons and exogenous transgenes. Additional mechanisms of piRNA action, namely target mRNA deadenylation, have also been reported, however, as they are implied in regulation of protein-coding gene expression rather than transposon silencing.

Piwi-mediated the transcriptional gene silencing (TGS) in which Piwi translocates to the nucleus to potentially interact with nascent transcript or DNA at the target locus, which in turn leads to heterochromatin formation and transcriptional repression as shown in figure.

In D. melanogaster, Piwi localises to the nucleus and initiates repressive histone H3K9 trimethylation and RNA polymerase II stalling. Whether Piwi interacts with the nascent transcript or directly with DNA is not understood. The zinc-finger protein Gtsf1 likely directly interacts with Piwi, whereas the heterochromatin protein Hp1 binds to H3K9me3. Mael acts downstream of H3K9me3 methylation and is required for POL II repression; however, its mechanism of action also remains to be determined. In parallel to transcriptional gene silencing (TGS), posttranscriptional gene silencing (PTGS; i.e. slicing) plays a well-defined role in D. melanogaster piRNA-mediated transposon silencing.

Molecular Techniques

• 1. Knockoutmice :

The technique of gene targeting allows for the introduction of engineered genetic mutations into a mouse at a determined genomic locus. The process of generating mouse models with targeted mutations was developed through both the discovery

• f homologous recombination and the isolation of murine embryonic stem cells

(ES cells). Homologous recombination is a DNA repair mechanism that is employed in gene targeting to insert a designed mutation into the homologous genetic locus. Targeted homologous recombination can be performed in murine ES cells through electroporation of a targeting construct. These ES cells are totipotent and, when injected into a mouse blastocyst, they can diffentiate into all cell types of a chimeric mouse. A chimeric mouse harboring cells derived from the targeted ES cell clone can then generate a whole mouse containing the desired targeted mutation. The initial step for the generation of a mouse with a targeted mutation is the construction of an efficient targeting vector that will be introduced into the ES cells.

The process of gene targeting provides a means to alter a specified gene in order to

better discern its biological role. Through homologous recombination, an engineered mutation can be directed to a designated genetic locus. In this manner, a potentially important genomic clone can directly be utilized to create a mutation into a selected gene. Even amongst the 2.5 Gb of the mouse genome, the cellular DNA repair mechanisms are able to align a targeting vector with its corresponding region of homology and cause recombination into the chromosome. The goal of transgenic technology is to overexpress a gene to study its biological role in vivo, homologous recombination is typically employed to create a ‘loss of function’ mutation. The most common application of gene targeting is to produce knockout mice, where a drug resistance marker replaces an essential coding region in a genetic locus. A gene inactivation is the best way to delineate the biological role of a protein and gene targeting is a direct means to disrupt a gene’s open reading frame and block its expression in a mouse.

The knockout mouse has been a valuable tool for geneticists to discern the role

• f a gene in embryonic development and in normal physiological homeostasis.

Mice act as a good analogue for most human biological processes since both species share about 99% of the same genes. Additionally, mice are useful experimental animals because they are small, have relatively short life spans, and are prolific. The targeted deletion of a gene in a mouse provides an important means to determine the biological role of a genetic allele. While useful to study in vivo gene function, some knockout mice have also additionally served as valuable animal models for human genetic diseases. The pharmaceutical companies obtain clues about inhibiting a protein by first looking at the phenotype of a knockout mouse. Thus, knockout mice can provide insight into a gene’s physiological role in humans.

Vector design

For recombination to occur at all in a cell, around 2 kb of sequence homology is

• required. However, 6 to 14 kb of homology is typical for targeting constructs.

In addition, linear DNA was found to be the preferred substrate for the recombination proteins. The actual targeted event takes place in only a small percentage of cells, as homologous recombination occurs about thousand fold lower than random

• insertions. During a stem cell experiment, only about 10−2 to 10−3 of the DNA

integrations are homologous recombination events. Therefore, a thorough screening process by Southern blot or by PCR is necessary to identify the cells with the targeted event. Electroporation became the preferred way to deliver the replacement vectors into a large number of stem cells. stem cells. While microinjection has a better targeting ratio (1:15 targeted recombinants to random integrants), a mass delivery system is needed to introduce the targeting constructs into cells and electroporation provided the most efficient technique (1:2,400 targeting ratio). However, since the transformation efficiency is low (10-

3), a positive selection marker is needed to enrich clones that have inserted the

targeting vector into their genome.

Primarily, two classes of vectors have been used to generate targeted mutations. Most gene targeting experiments employ replacement vectors as shown in figure, which have been particularly instrumental for efficiently generating knockout mice. In the design of a replacement vector, the open reading frame of a genomic clone is disrupted by the placement of an intervening drug selection marker.

Two homologous recombination events function to insert the targeting construct containing the drug resistance gene into a homologous genetic locus.

Gene inactivation through a replacement vector: Homologous recombination with a replacement vector requires a positive selection marker (neor), a negative selection marker (HSV-tk), and two targeting arms.

The drug resistance gene works for the positive selection of cells that have integrated the targeting vector into their chromosome. Neomycin is the most common drug used for positive selection. Integration of the neomycin phosphotransferase (neo-r) gene allows for resistance to neomycin, an aminoglycoside that interferes with protein synthesis in eukaryotic cells. Other drug selection markers include resistance genes for puromycin and hygromycin. The additional placement of the HSV thymidine kinase (HSV-tk) gene adjacent to

• ne of the vectors arms containing targeted homology to a genetic locus. Random

integrants will usually contain an intact of the HSV-tk gene when inserted into the

• genome. Cells with random integrants are killed during negative selection

through treatment with gancylovir that require phosphorylation by HSV-tk to inhibit DNA synthesis. Insertion of the neo-r gene is selected for by treatment of cells with neomycin sulfate (G418) in tissue culture. The negative selection marker (HSV-tk) is not recombined into the chromosome and is lost during gene

• targeting. If the targeting construct is randomly integrated anywhere in the

genome, the HSV-tk gene would be intact. Random integrants can be selected against via gancyclovir or FIAU treatment.

• 2. Phage Display Technique :

The Phage display technique is one of the most effective molecular diversity techniques is phage display. This technology is based on a direct linkage between phage phenotype and its encapsulated genotype, which leads to the presentation of molecule libraries on the phage surface. The phage display is utilized in studying protein ligand interactions, receptor binding sites and in improving or modifying the affinity of proteins for their binding partners. Generating monoclonal antibodies and improving their affinity, cloning antibodies from unstable hybridoma cells and identifying epitopes, mimotopes and functional or accessible sites from antigens are also important advantages of this technology. Techniques originating from phage display have been applied to transfusion medicine, neurological disorders, mapping vascular addresses and tissue homing

• f peptides.

Phages have been applicable to immunization therapies, which may lead to development of new tools used for treating autoimmune and cancer diseases.

Since then, a large number of phage displayed peptide and protein libraries have been constructed, leading to various techniques for screening such libraries. This technology has had a major influence on the work and discoveries done in the fields

• f immunology, cell biology, pharmacology and drug discovery.

The phage display allows the presentation of large peptide and protein libraries

• n the surface of filamentous phage, which leads to the selection of peptides and

proteins, including antibodies, with high affinity and specificity to almost any target. The technology involves the introduction of exogenous peptide sequences into a location in the genome of the phage capsid proteins. The encoded peptides are expressed or “displayed” on the phage surface as a fusion product with one of the phage coat proteins. This way, instead of having to genetically engineer different proteins or peptides one at a time and then express, purify, and analyze each variant, phage display libraries containing up to

• E. coli filamentous bacteriophages (f1, fd, M13) are commonly used for phage
• display. Most antibodies and peptides are displayed at phage proteins pIII and
• pVIII. The major coat protein (pVIII) is a product of gene 8 expression and occurs

in nearly 3000 copies, therefore it is used to enhance detection signal when phage displayed antibody associates with antigen. Moreover modifications of pVIII are made to increase the efficiency of display onto pVIII. In comparison, a minor coat protein (pIII) consists of 406 amino acid residues and occurs at the phage tip in 3 to 5 copies. The vast majority of peptides and folded proteins are displayed as fusions with pIII protein.

PIII, pVI, pVII, pVIII and pXIX represent phage proteins. Exogenous peptides are expressed or “displayed” usually on pIII or pVIII. The loss of coat protein functionality was the major limitation of the phage display technology, however this problem was overcome by hybrid phages and coat protein modifications. These virions consist of the complete wild type genome and a copy of fusion gene which might occur as an insert in phage genome or as phagemid a vector that contains the

• rigins of replication for phage and its host, gene 3 with

appropriate cloning sites and an antibiotic-resistance gene. Moreover, the phagemid encoding polypeptide-pIII fusion requires hybrid with helper phage for packing into M13 particle.

The helper phage contains a slightly defective origin of replication (e.g. M13KO7) and supplies all the structural proteins required for generating a complete virion. Thus, both wild pIII protein and polypeptide pIII fusion protein will be present

• n the phage surface.

The ratio of polypeptide-pIII fusion protein to wild type pIII may range between 1 to 9 and 1 to 1,000 depending on the type of phagemid, growth conditions, the nature of the polypeptide fused to pIII and proteolytic cleavage of antibody-pIII fusions. Moreover, hybrid phage system enables displaying large proteins with all five M13 coat proteins as N-terminal fusions with pIII, pVIII, pVII and pIX and also as C-terminal fusions with pVI, pIII, and pVIII. Due to the naturally occurring translational stop codon in the 3'-region of reverse transcribed mRNAs in M13 display system, expression of cDNA libraries could be difficult.

The usage of T7 is an alternative for M13 display. The T7 phage display system has been widely used, due to its extreme robustness and stability in conditions that inactivate other phages.

The phage T4 HOC/SOC bipartite display system could be applied to cDNA

• expression. It displays larger proteins in high copy number and inserts with stop

codon on the C-terminal of SOC (small outer capsid) protein that occurs in 810 copies

• r N-terminal of HOC (highly antigenic outer capsid) protein that
• ccurs in 155 copies.

The Phage lambda is capable of displaying complicated, high molecular mass proteins as fusions with N- or C-terminal of pD head protein that occurs in 405 copies or C-terminal of pV tail protein that occurs in 6 copies.

The structure of most antibodies consists of two heavy chains (variable (VH), diversity (D), joining (JH), and constant (CH) region) and a pair of light chains (variable (VL), joining (JL) and constant (CL) region) linked by non-covalent bonds and disulfide bridges. The phage display technology has provided the ability to create antibody libraries that contain a great number of phage particles, from which each one encodes and displays different molecules. Therefore biopanning—the procedure of specific binders selection—is essential for enriching the desired molecule level. The biopanning method is based on repeated cycles of incubation, washing, amplification and reselection of bound phage. The target molecule is immobilized

• n solid support as microtiter plate wells, PVDF membrane, column matrix or

immunotubes, magnetic beads and even on whole cells. The several rounds of selection cycle are necessary to achieve the desired binding activity of obtained monoclonal phage antibodies. For determining this activity several tests are used, for example enzyme-linked immunosorbent assay (ELISA), fluorometric microvolume assay technology (FMAT) or chromophore-assisted laser inactivation (CALI). The type of solid support, time of binding and washing as well as antigen concentration have affected the level of selection.

A proper design of the biopanning procedure enables the selection of antibodies to unique epitopes

3.Expressed sequence tags (ESTs)

Expressed sequence tags (ESTs) are single-pass reads of approximately 200– 800 base pairs (bp) generated from randomly selected cDNA clones. Since they represent the expressed portion of a genome, ESTs have proven to be extremely useful for purposes of gene identification and verification of gene predictions. The use of expressed sequence tags dates back to the early 1980s, when Putney and colleagues sequenced inserts from 178 clones derived from a rabbit muscle cDNA library. Today ESTs are still being exploited for purposes of novel gene discovery, especially for those organisms for which economies of scale preclude complete genome sequencing. The advent

• f

cheaper and faster sequencing technologies has generated an exponential growth in the number of ESTs that have been generated.

The process of generating ESTs starts with the initial purification of pools of mRNAs from either a whole organism or specific tissues.

(A)Depending upon the size of the organism, mRNA may be collected either from the whole

• rganism or from specific tissues. (B) Pools of

mRNA are extracted and purified (typically on the basis of their poly-adenylation). (C) A cDNA library is then constructed from this pool and clones are randomly picked for a single-pass sequencing read (D). The raw trace data are then processed to derive the underlying sequence (E).Further processing identifies and removes low-quality sequence and contaminating sequence associated with the cloning and/or sequencing vectors. The purified sequence data can then be submitted to the international repository of ESTs – dbEST – along with associated metadata detailing the source of ESTs and methods used in their generation. (F)Finally, ESTs may undergo postprocessing, for example, they may be clustered on the basis of sequence similarity to derive groups of sequences that putatively derive from the same gene. Consensus sequences derived from these clusters may be further annotated, e.g. via BLAST searches and used to construct queryable databases such as Parti GeneDB.

The mRNAs are isolated on the basis of their 30 poly-A tails and reverse transcribed to create libraries of cDNAs cloned into an appropriate vector. Individual clones from these libraries are then selected (typically at random) and subjected to a single sequencing reaction using universal primers that can be associated with either end of the insert. Downstream of the sequencing reaction, a number of sophisticated bioinformatics pipelines have been developed that process the raw sequence read to remove low- quality sequence information and contaminating vector sequence. The length of the generated read varies according to the sequencing technology used. For example, the latest capillary-based technologies can produce reads of up to 800 base pairs of relatively high quality. The resultant high-quality trimmed sequence is then deposited in the specialized dbEST database. At each stage of this process, a number of strategic decisions must be made according to the objectives of the EST project. These include the source and number of clone libraries to generate ESTs, the sequencing technology itself and the software to employ for sequence processing.

Generation of c-DNA libraries

The choice of material and method for library production is dependent, to some extent, on the aims of the project. For example, projects attempting to identify new stress genes in plants would likely use material from plants that have been exposed to a particular stress. Considerations for material include life cycle stage, sex and tissue types. Even for relatively small organisms or cell- or tissue-types within larger organisms, techniques such as laser microdissection microscopy offer avenues for tissue- specific cDNA library generation. It should be noted that while continued sequencing from the same individual library may be useful for determining the relative abundance of transcripts, it does lead to diminishing returns in terms of novel gene discovery. Projects focused on gene discovery should therefore consider the generation and sampling of multiple libraries. Over the past 5 years, we have seen significant improvements in DNA sequencing resulting in an increase in both the length and the quality of reads. Two recent breakthroughs in sequencing have been the successful deployment of ‘‘massively parallel’’ platforms based on reversible dye termination sequencing and pyrosequencing.

With the increasing volume of sequences now being generated, bioinformatics pipelines are now becoming essential tools for their processing. The main repository for EST sequence data is dbEST available through the National Center for Biotechnology Information (NCBI) Web portal (and the Web portals

• f the other members of the International Nucleotide Sequence Database

Consortium). The NCBI database is readily queryable through the Entrez nucleotide or BLAST interfaces. In addition, it is now possible to upload and retrieve the raw trace (or sequencing chromatogram) information associated with individual

• ESTs. This latter feature allows users to perform their own analyses to detect low-

quality or vector-contaminating sequence rather than relying on the submitters to correctly identify such sequence. ESTs have proved to be a cheap and effective tool for gene discovery across a variety of scales. For example, applied to parasites such as the sheep scab mite Psoroptes ovis, the generation of even a few hundred ESTs successfully identified a number of possible antigens that are now the focus of more in-depth functional investigations.

The increasing number of ESTs from a diverse set of organisms deposited in dbEST provides an unrivalled opportunity to explore eukaryotic diversity in a depth not possible using only the few fully sequenced genomes. The study of expression patterns of genes across a range of cell types, life cycle stages, or environmental conditions has proved to be extremely useful for understanding gene functions. A major technology in the field of proteomics is tandem mass spectrometry, which attempts to match peptide fragments to known protein sequences. However, even for well-studied organisms, the limited number of such sequences in protein databases such as Swiss-Prot results in a computational bias against poorly characterized proteins. Thus, due to their coverage, ESTs are beginning to find widespread appeal and also allow the identification of peptides from alternative spliced isoforms.

4.Yeast two-hybrid

The yeast two-hybrid system is a powerful and commonly used genetic tool to investigate interactions between artificial fusion proteins inside the nucleus of yeast. The yeast two-hybrid system (Y2H) was first developed in 1989 and revolutionized the process of searching for and identifying interacting proteins. To date, the Y2H system has proven to be a useful and sensitive method for detecting not only stable interacting proteins but also weak and transient protein interactions. Because Y2H is performed in vivo, the great advantage of the system is that the testing proteins are more likely to be in their native conformations, which may lead to increased sensitivity and accuracy of detection. Importantly, the Y2H system is complementary to biochemical methods such as co- immunoprecipitation/pulldown followed by western blotting

• r

mass spectrometry analysis to increase accuracy and dynamics for a more complete and reliable map of interactions.

Notably, the Y2H method has been modified and improved greatly in recent years, including applications in protein–DNA interactions and yeast three-hybrid, and has proven to be amenable to interaction studies of membrane proteins, DNA binding proteins, and RNA binding proteins. The Y2H technique allows detection of interacting proteins in living yeast cells Interaction between two proteins, called bait and prey, activates reporter genes that enable growth on specific media or a color reaction. Y2H can be easily used for studies of protein interactions on a genome-wide scale, as shown for viruses like bacteriophage T7, Saccharomyces cerevisiae, Drosophila melanogaster, Caenorhabditis elegans and humans. Experimental Y2H data have been a crucial part in establishing large synthetic human interactomes or to dissect mechanisms in human disease.

Using the Matchmaker GAL4-based Y2H system as an example, the principle

• f the Y2H system is illustrated in the following Figure

Bait proteins are expressed as a fusion to the GAL4 DNA binding domain (DNA-BD), while prey proteins are expressed as fusions to the GAL4 activation domain (AD). When bait and prey fusion proteins interact in yeast nucleus, the DNA-BD and AD are brought into proximity to restore to a functional GAL4 transcriptional activator, which binds onto an upstream activating sequence (UAS) of reporter genes for transcriptional activation. The Y2H system has been widely used to detect the interactions of a wide range of proteins from yeast, bacteria, animal, and plant systems.

In A. tumefaciens, we showed the interaction of the T6SS sheath components TssB and TssC41 by Y2H assay. For the Y2H assay, each bait and prey plasmid pair was co-transformed into the Saccharomyces cerevisiae strain AH109. The transformants were selected by their growth on synthetic dextrose (SD) minimal medium lacking tryptophan (Trp) and leucine (Leu) (SD-WL medium). The positive interaction of expressed fusion proteins was then determined by their growth on SD lacking Trp, Leu, adenine (Ade), and histidine (His) (SD-WLHA) medium at 30 °C for at least 3 days. The positive interactions were observed only for plasmid pairs expressing TssB and TssC41 but not when each of them co-expressed with vector only, suggesting the specific interactions of TssB and TssC41.

TssB and TssC41 interact with each other in yeast strain AH109. SD-WL medium (SD minimal medium lacking Trp and Leu) was used for the selection of plasmids. SD-WLHA medium (SD minimal medium lacking Trp, Leu, His, and Ade) was used for the auxotrophic selection of bait and prey protein interactions. The positive interaction was determined by the growth

• n

SD-WLHA medium at 30 °C for at least 2 days

5.Yeast three-hybrid

The yeast three-hybrid system has become a useful tool in analyzing RNA–protein

• interactions. The RNA–protein interactions underlie diverse biological processes,

from pattern formation to the replication of viruses. As a result, methods have been developed to analyze RNA– protein interactions using molecular genetics. A yeast three-hybrid system has enabled the identification of naturally occurring RNA and protein partners, and the dissection of higher-order RNA–protein

• complexes. In the yeast three-hybrid system, a chimeric protein containing both a

DNA- and RNA-binding domain tethers an RNA to the promoter of a reporter gene Typically, this protein consists

• f

a LexA/MS2 coat protein fusion. A hybrid RNA binds to the MS2 portion via tandem MS2-binding sites The RNA also contains a sequence of interest, X, which binds to an RNA-binding polypeptide, Y. Y is linked to a transcription activation domain (AD). When the requisite interactions occur, the reporter gene is activated. Typically, the interaction between RNA X and protein Y is monitored by assaying HIS3 and LacZ expression levels.

The expression level of lacZ gene can be determined in vitro by measuring the β- galactosidase activity, or visualized in vivo by plating the yeast transformants on media supplemented with X-Gal. On the other hand, HIS3 is the gene encoding imidazole glycerol-phosphate dehydratase (His3p) and its expression confers the ability to grow on a medium lacking histidine. 3-amino-1,2,4-triazole (3-AT) is a competitive inhibitor of HIS3 gene product, and therefore cells containing more His3 product can survive at higher concentrations of 3-AT in the medium. Thus, the level of 3-AT resistance of the yeast cells reflects the HIS3 expression level and consequently the strength of the RNA–protein interaction in the yeast three-hybrid context. Qualitatively, weak and strong interactions can be discriminated phenotypically using HIS3. This feature has been used to isolate mutations in the RNA or protein that alter affinity. However, a systematic examination of the relationship between in vitro affinity of an RNA–protein interaction and reporter gene activity has not been reported. This limits interpretation of the data, and hampers the design of new applications.

• 6. Activity Gel Assay

Several proteins are remain uncharacterized in terms of their functions in health and disease states, annotation of protein function is an important task in current biological and medicinal study. Even if we limit our consideration to enzymes, it remains the case that connecting enzymatic functions to changes in metabolites, in

• ther words, annotation of enzymes responsible for certain enzymatic reactions,

can yield huge progress in understanding the roles of proteins in living systems. For example, the discovery of the physiological substrate of dipeptidyl peptidase IV (DPPIV) made the enzyme a promising target for treatment of diabetes. However, despite the great potential value of such studies, there has been little progress in methods to annotate proteins on the basis of their biochemical activities. A conventional approach for this purpose would be column-chromatography- based proteome separation coupled with enzymatic assays. However, this is slow and requires large amounts of sample, and clear-cut separation is often difficult. Since plural enzymes may exhibit a particular activity in physiological or pathological states, it is important to identify them in a comprehensive manner to select potential targets for drug development.

Therefore, we need a method for sensitive, reliable, rapid, and comprehensive detection of proteins with specified activities. One platform for this purpose is zymography in which the proteome is separated by nondenatured polyacrylamide gel electrophoresis (PAGE) and colorimetric or fluorometric assays are performed on the gel to visualize protein spots with desired activities. The method requires only small amounts of sample and gives sharp separation, and two dimensional electrophoresis can give a comprehensive “activity map”. However, the sensitivity is low, especially with colorimetric substrates. For example, the detection limit of β-galactosidase activity with the colorimetric substrate X-gal is only 100 ng. Fluorescent substrates would increase the sensitivity, but in the conventional system, the fluorescent product readily diffuses in the gel, making it difficult to determine the precise location of the target protein.

Effective way to overcome the problem of diffusion by dicing the electrophoresis gel into small pieces that are separately loaded into wells of multiwell plates. The fluorometric assay can then easily identify wells containing active proteins This assay is called diced electrophoresis gel (DEG) assay

The method not only enables fluorescent substrates to be used in zymography to achieve much higher sensitivity but also allows the use of various detection platforms, including LC−MS-based analysis. Tested the feasibility of the DEG assay by detecting β-galactosidase on the gel after blue native PAGE with the fluorescent substrate TG β-Gal. The usefulness of the method was confirmed by detection of two glycosidases (β-glucosidase and β-galactosidase), two esterases (liver esterase and alkaline phosphatase), and two peptidases (leucine aminopeptidase and elastase) with appropriate fluorescent substrates. The detection limits ranged from 0.1 to 10 n

• 7. Helicase Assay

Helicases are a central class of enzymes found in all known organisms. By a combination of biochemical and genetic approaches, their activity has been shown to affect most metabolic processes that rely on nucleic acid unwinding in the cell, including DNA replication, transcription, translation, DNA repair

• r recombination.

At the molecular level, helicases are motor proteins that move directionally along a nucleic acid phosphodiester backbone and are able to separate annealed nucleic acid strands using energy derived from adenosine triphosphate (ATP) hydrolysis. Although the majority of the known helicase enzymes are involved in the process of unwinding duplex DNA or RNA. Some of these proteins have been known to act on less canonical substrates like protein-nucleic acids complexes or secondary DNA structures, in particular G-quadruplexes.

The most common assay for measuring helicase activity in vitro employs gel electrophoresis. The helicase activity gel : The DNA helicase activity gel system is analogous to a widespread assay for the detection of purified DNA helicase activities . In this assay, enzymatic displacement of a radioactive fragment from a non-radioactive single-stranded M13 DNA is detected by measuring an increase in the electrophoretic mobility of the displaced DNA versus the radioactive substrate. This assay works well with purified helicases but it cannot be applied to unfractionated crude extracts because nucleases degrade the substrate and products.

The activity gel assay is designed to avoid this problem by electrophoretically fractionating the proteins of a crude extract in an SDS-polyacrylamide gel. The fractionated proteins are then renatured and allowed to react with a partially duplex radioactive DNA substrate that is immobilized in the gel. The circular DNA substrates do not migrate during the electrophoresis of the sample because they presumably are interlocked with the network of polyacrylamide strands. Following electrophoresis the gel is washed extensively to remove the SDS and to promote the renaturation of the fractionated proteins. Next, the polyacrylamide gel is incubated in a reaction buffer allowing DNA metabolic activities in the samples to convert the DNA substrate into a form that subsequently can be blotted from the gel to a nylon filter. The resulting activity bands can be detected by subjecting the filter to autoradiography.

Although the activity gel assay was designed to allow detection of DNA helicases, the assay allowed the simultaneous detection of endonuclease and exonuclease

• activities. Electrophoresis has the obvious advantage to allow the determination of

the diversity and relative abundance of the molecular species present in the reaction. This is however a relatively cumbersome and low-throughput technique. Several additional methods based on fluorescence assays have been developed to overcome some of the limitations of electrophoresis. To improve tools aiming at characterizing the activity of DNA helicases towards G4 structures, recently developed a fluorescence-based helicase assay that allows testing the unwinding

• f different G4 structures by a helicase in high throughput and in real time.

Real-time helicase assay : Fluorescence is a sensitive technique that has the clear advantage of monitoring are action process in real time, at low concentration and at high throughput. For this reason, the helicase assay is designed to monitor the unwinding process of a fluorophore-labelled DNA system. Pif1 helicases translocate in a 5’to 3’ direction and require a 5’ tail to load on their substrate

Thus, the designed DNA system to contain this recognition element. The substrate has a single-stranded region, a G4 region, a duplex region and a 3’ tail

• f 17 nucleotides covalently attached on its 3’ end to a dabcyl residue. A

complementary oligonucleotide labelled at its 5’end with the FAM fluorophore is annealed to 15 nucleotides. When duplex is formed, the fluorescence of the FAM (orange symbol) labelling the single strand is quenched. In the first step of the reaction, the Pif1p helicase binds to the 5’ single-strand end. After ATP addition, the helicase proceeds in a 5’ to 3 direction unwinding first the G- quadruplex structure and then the duplex DNA. Unwinding of the duplex releases the labelled single-stranded probe, which emits a strong fluorescence as it is no longer quenched by dabcyl (purple symbol) on the complementary strand.

8.Chromatin Immuno-precipitation (ChIP) Assay

Techniques commonly used to characterize protein–DNA interactions include electrophoretic mobility shift assay (EMSA), DNAse footprinting and chromatin immunoprecipitation (ChIP) assays. Though, EMSA is a rapid and sensitive method to detect protein–DNA interactions, despite its advantages it has limitations like the samples are not in chemical equilibrium during the electrophoresis step and also many complexes are significantly more stable in the gel than at free solution. ChIP assays minimize these limitations and provide an efficient tool to determine these protein–DNA interactions occurring in vivo, by immunoprecipitating chromatin with specific antibodies. The ChIP assay has been adopted as a powerful method for the analysis of proteins interacting within a native chromatin environment and is versatile enough for adaptation for a variety of purposes. This assay has been utilized in yeast, drosophila, tetrahymena, Caenorhabditis elegans, various mammalian cell lines, and even on whole mouse embryos for the analysis of low abundance transcription factor binding.

The ChIP assay is a multistep process and each step needs to be standardized for

• btaining optimum results. In this technique, intact cells are treated with

formaldehyde to covalently link protein to DNA (X-ChIP), the nucleoprotein complexes are then sheared either mechanically or by enzymatic digestion. The resultant soluble cross-linked DNA-protein complexes enriched by immunoprecipitation.

The retrieved complexes are then analyzed by PCR amplification with gene- specific primers to detect and quantify specific DNA targets. The main drawback of ChIP is its inherent variability. This may arise due to variabilities in the cross-linking, immunoprecipitation, and protein–DNA washing

• efficiencies. Stringent control over experimental conditions may reduce these

variabilities. ChIP applications for discovering novel genes dependent on specific transcription

• factors. The ChIPed DNA can be ligated

and cloned into vector. Colonies having inserts are identified by restriction enzyme digestion using enzymes in the

• polylinker. Plasmids with inserts greater

than 500 bp are studied.

• 9. Designing probe

A stretch of DNA or RNA sequence that can detect a target sequence in the genome is called as probe. Molecular probes are small DNA or RNA segments that recognize complementary sequences in DNA or RNA molecules that allow identification and isolation of these specific sequences from an organism. These molecular markers have acted as versatile tools and have found their own position in various fields like taxonomy, physiology, embryology and genetic engineering. Although, initially these probes were developed and used for genetic engineering research but are now frequently used for a variety of purposes including diagnosis of infectious diseases, identification

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food contaminants, variety

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microbiological tests and forensic tests. Probes can also be used to identify different varieties of crop species. For basic studies in molecular biology laboratories these are frequently used for identification and isolation of genes or related sequences. In practice, double and single standard DNAs, mRNAs, and other RNAs synthesized in vitro are all used as probes. DNA/ RNA probe assays are faster and sensitive so that many conventional diagnostic tests for viruses and bacteria involving culturing of the organisms are being fast replaced by molecular probe assays.

Molecular probes can be broadly categorized into DNA probes and RNA probes, sometimes cDNA probes and synthetic oligonucleotide probes can also be used for various purposes. Preparation of Probes Different types of probes can be prepared in various ways.

DNA Probes : Extract the DNA from an animal or plant tissue. Digest

extracted DNA with a restriction enzyme such as EcoRI or Hind III which cuts DNA at specific sites or positions where a specific sequences recognized by the enzyme is found. Run the digested DNA on an agarose or polyacrylmide gel electrophoresis to separate fragments of different sizes. Isolate DNA of specific fragment from a particular band identified through southern blots by hybridization with specific labeled mRNA or cDNA molecules. Clone this DNA in a vector. Allow chimeric vector to infect bacteria for multiplication where it can make billions of copies. DNA probes prepared in this manner can be used for southern blotting and Restriction fragment length polymorphism (RFLP) analysis.

RNA Probes : High specific activity RNA probes or riboprobes may also be

synthesized from DNA templates cloned in expression vectors such as SP6 (which infects Salmonella typhimurium) and T7 phage (infects E.coli). This is achieved through RNA synthesized in vitro and labeled simultaneously with labeled nucleotides.

The probes should be transcribed into uniform lengths, it is practice to linearize the plasmid by cleaving it with a restriction enzyme. The vector thus carries the input in the following order phage promoter-Enzyme 1- Enzyme 2. The template DNA is inserted at the number 1 site and composite is treated with restriction enzyme 2. So now this creates a linear DNA with a promoter, the template DNA and automatic termination site at the end cleaved with enzyme 2. The mRNAs fall of when they reach this end of the vector. Such templates reflect to run of templates and are uniform in size and are easier to isolate from the reaction mixture. RNA probes prepared in this manner can be used for northern blotting and in situ hybridization.

cDNA Probes : A DNA sequence corresponding to a part of a specific gene can

be obtained by reverse transcription of mRNA. cDNA thus obtained can be cloned and used as a probe.

Synthetic oligonucleotides as probes : DNA probes with known nucleotide

sequence can also be synthesized chemically using automated DNA synthesizers. These synthetic probes will be efficient only when they are not more than 2040 nucleotides in length.

Labeling of probes :

The detection of homologous sequences after hybridization with the probe is like finding a needle in the hay-stock. Therefore, for the success of DNA probe assay it is necessary to develop simple, safe and sensitive techniques for their use. As probes transmit no signal of their own they have to be either labeled with radioactive isotopes or coupling of non-radioactive signal molecules to the probes without impairing the hybridization ability of these

• probes. These signal molecules may include fluorescent antibodies, enzymes that

produce color changes in dyes and chemiluminescent catalysts.

Methods for Labeling of Probes There are two methods for labeling of

• probes. i.e. end labeling and nick translation.

End labeling : In this technique probe is isolated and end labeled by removing the 5'-terminal phosphate using alkaline phosphatase first and adding a 32P-labeled phosphate with the help of a kinase. Nick Translation: It is one of the commonly used techniques for producing a radioactive probe. A purified phage or plasmid vector containing a cloned genomic or cDNA sequence is treated with a small amount of pancreatic DNase which hydrolyzes the phosphodiester bonds between nucleotides. At very low concentration the DNase produces only scattered “nicks” in one or other strand of the duplex DNA.

DNA polymerase and radioactively labeled deoxynucleotides are also added to the DNA sample. Using the unharmed strand as template, the DNA polymerase synthesizes a new second strand using exposed 3' end at a nick site as primer, which then displaces the existing DNA from the 5’ end of the nick. Radioactive nucleotides are incorporated into the new strand, so, a single standard probe is created when the duplex DNA is denatured.

10.Epitope Tagging

Epitope tagging is a method of expressing proteins whereby an epitope for a specific monoclonal antibody is fused to a target protein using recombinant DNA techniques. By choosing an epitope for which an antibody is available, the technique makes it possible to detect proteins for which no antibody is available. This is especially useful for the characterization of newly discovered proteins and proteins of low immunogenicity. By selection of the appropriate epitope and antibody pair, it is possible to find a combination with properties that are suitable for the desired experimental application, such as Western blot analysis, immunoprecipitation, immunochemistry, and affinity purification. The fusion gene is cloned into an appropriate expression vector for the experimental cell type and host cells are transfected. The fusion protein can then be detected and/or purified using a monoclonal antibody specific for the epitope tag.

Selection of an epitope tag and monoclonal antibody will depend upon the

• experiment. The more widely used epitope tags are the HA, myc and FLAG tags.

Monoclonal antibodies directed against these epitope tags are readily available. A primary factor in choosing a tag to use is the availability of an antibody. It is advisable to perform control experiments to demonstrate that the antibody does not recognize non-tagged proteins within the cell. The use of the FLAG tag, which consists of the eight-amino-acid sequence N-Asp-Tyr-Lys-Asp-Asp-Asp- AspLys-C. A number of antibodies and antibody conjugates to the FLAG tag are available for use in applications such as western blotting, immunostaining, and immunoaffinity purification. The gene encoding the protein of interest is fused to the FLAG sequence by cloning into an appropriate vector and then expressed in host cells. The FLAG tag is most commonly placed at the N-terminus or C- terminus of the protein. In N-terminal expression vectors the FLAG tag is preceded by either a secretion signal sequence or a translational initiation

• codon. Insertion of a target gene into an appropriate expression vector is best

accomplished by polymerase chain reaction (PCR) in order to achieve the correct translational phasing.

Moreover, the 24-bp FLAG coding sequence can be incorporated into

• ligonucleotide primers for creation of FLAG fusions by PCR in any suitable
• vector. The presence of the proper DNA coding sequence at the junction of the

inserted gene and the FLAG sequence should be confirmed by DNA sequencing prior to transfection of host cells. The single most important application

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epitope-tagging is co-

• immunoprecipitation. In this technique, a bait protein fused to an epitope tag is

transfected into cells. Following lysis of the transfected cells, the expressed bait protein is immunoprecipitated with anti-tag antibody coupled to beads. Proteins that interact directly with the bait protein are co-immunoprecipated. Typically, the protein complex is then analyzed by SDS-PAGE. A common variation of this method uses co-transfection of the bait and interacting proteins, each tagged with a different epitope tag. After co-immunopreciptiation using antibody to the epitope-tagged bait protein, the interacting protein may be detected by Western blot analysis using antibody against the other epitope tag. The power of this technique has contributed greatly to our understanding of protein interaction networks.

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