the use of oligonucloutide tool in DNA

 the use of oligonucleotide tool in BIOLOGY

 

The use of oligonucleotides as tool in cell biology

Oligonucleotides, or oligos, are short single strands of synthetic DNA or RNA that serve as the starting point for many molecular biology and synthetic biology applications! From genetic testing to forensic research and next-generation sequencing, an oligo may very well be the starting point. (1).

How are oligos made?

Custom DNA oligos are made by a process called synthesis or more specifically, solid-phase chemical synthesis. This is a method in which the 4 nucleic acids, A, T, C, and G, are added one by one to form a growing chain of nucleotides. They are built on an oligo building block called a phosphoramidite. During these cycles of adding one nucleotide or base to another, the chain grows in the 3’ to 5’ direction. At the end of synthesis, or all the cycles adding each base, the result is a full-length oligo (1). After the oligo is completed, it typically gets desalted or sometimes even purified. Desalting an oligo removes the salts used in the synthesis process. After this step, the oligo is ready to use for applications like PCR (1).

However, during synthesis there are also shorter chains or failure sequences that form as well. This is because no chemical reaction is 100%, so each time a base is added, it can fail to attach and may help form a smaller side chain. These smaller side chains or failure sequences can compete with the full- length sequence in some downstream applications and can potentially impact results. If the downstream application requires only the full-length sequence, then there are several other purification options that create a purer oligo by removing these failure sequences. These include HPLC, Cartridge or PAGE. Applications like NGS (Next-Generation Sequencing) or mutagenesis require very pure oligos. (1).

What are oligos used for?

The most common example of an application that oligos are known for is PCR or polymerase chain reaction. PCR is the technique of making many copies of a fragment or strand of DNA to then generate thousands or millions of more copies for use in other downstream applications like cloning or sequencing. Researchers use oligos that are anywhere between 20-35 bases long called primers to start copying or amplifying. This DNA primer is usually custom designed to match the target sequence of DNA for copying. The ability for researchers to design their own custom DNA oligos for their experiments has opened new doors in fields like Molecular Biology and Synthetic Biology. (1).

Antisense oligonucleotides provide a promising approach to investigating gene function in vivo; buttheir ability to offer unambiguous insights into phenotypes has been debated. The recent use ofmorpholino antisense oligonucleotides in zebrafish embryos may prove a major advance, but rigorouscontrols are essential. (2).

 

 

The escalating pace at which genome sequencing projectsare completed has increased the need for high-throughputmethods for controlling gene expression. One possibleapproach is the use of antisense oligonucleotides to bindmRNA and prevent protein synthesis. In theory, this strategyallows rapid progress from synthesis of oligomers to obser-vation of phenotype [3]. In practice, antisense technologyhas been plagued by a propensity for nonspecific interac-tions, and these have slowed its wide application to biologi-cal investigations [3]. Recently, however, improvements inthe chemical properties of oligonucleotides and in ourunderstanding of their mechanism of action have combinedto make their successful use more likely. Work from anumber of laboratories now suggests that morpholinooligonucleotides can be microinjected into zebrafish [4-8], sea urchin [9] or Xenopusembryos [10], where they blockgene expression and produce phenotypic effects during theearly stages of development.

Morpholino oligonucleotides

Morpholino oligonucleotides are nonionic DNA analogsavailable from Gene Tools LLC [11, 12]. They possess alteredbackbone linkages compared with DNA or RNA (figure 1).In spite of their altered backbone, morpholinos bind to com-plementary nucleic acid sequences by Watson-Crick base-pairing. This binding is no tighter than binding of analogousDNA and RNA oligomers, necessitating the use of relativelylong 25-base morpholinos for antisense gene inhibition. Thebackbone makes morpholinos resistant to digestion by nucleases. Also, because the backbone lacks negative charge,it is thought that morpholinos are less likely to interact non-selectively with cellular proteins; such interactions often obscure the observation of informative phenotypes. Thestrengths of morpholinos as tools for investigating verte-brate development are well described in a recent review byEkker [6]. Their greatest advantage is that phenotypes canbe rapidly observed in B0 animals using a relatively inexpen-sive method.

 

“aptamer”

The terms “aptamer” and “SELEX” were introduced by two independent groups in 1990. (13, 14), the term “aptamer” refers to small nucleic acid ligands that exhibit specific therapeutic functions and an unambiguous binding affinity for their targets. Conversely, Systematic Evolution of Ligands by EXponential enrichment (SELEX) technology is the method used for aptamer development. Although using small molecule nucleic acids as therapeutics has been explored for decades, development of SELEX and aptamer technology revolutionized this field.

The most important property of an aptamer, from the Latin aptus (to fit), is its high target selectivity. These short, chemically synthesized, single-stranded (ss) RNA or DNA oligonucleotides fold into specific three-dimensional (3D) structures with dissociation constants usually in the pico- to nano-molar range(15). Moreover, in contrast to other nucleic acid molecular probes, aptamers interact with and bind to their targets through structural recognition (Figure 2), a process similar to that of an antigen-antibody reaction. Thus, aptamers are also referred to as “chemical antibodies.”

Figure 2

 

Schematic diagram of aptamer binding to its target

Due to their small size and oligonucleotide properties, aptamers offer several advantages over protein antibodies in both their extensive clinical applicability and a less challenging industrial synthesis process. Specifically, (i) aptamers can penetrate tissues faster and more efficiently due to their significantly lower molecular weight (8–25 kDa aptamers versus ~150 kDa of antibodies). Therefore, aptamers penetrate tissues barriers and reach their target sites in vivo more efficiently than the larger-sized protein antibodies. (ii) Aptamers are virtually nonimmunogenic in vivo. In principal, as aptamers are oligonucleotides they should not be recognized by the immune system. In practice, a recent clinical study showed that aptamers did not stimulate an immune response in vivo (16, 17), as compared to protein antibodies that are highly immunogenic, especially following repeat injections. (iii) Aptamers are thermally stable. Based on the intrinsic property of oligonucleotides, even after a 95 °C denaturation, aptamers can refold into their correct 3D conformations once cooled to room temperature. In comparison, protein-based antibodies permanently lose their activity at high temperatures. More importantly, a well-established synthesis protocol and chemical modification technology lead to (iv) rapid, large-scale aptamer synthesis and modification capacity that includes a variety of functional moieties; (v) low structural variation during chemical synthesis; and (vi) have lower production costs. Moreover, aptamers specifically recognize a wide range of targets, such as ions, drugs, toxins, peptides, proteins, viruses, bacteria, cells, and even tissues (18, 19, 20, and 21). In the clinic, aptamer-based therapeutics is gaining momentum. For example, Macugen, a modified RNA aptamer, specifically targets vascular endothelial growth factor. It has been approved by the US Food and Drug Administration (FDA(26)) for the treatment of wet age-related macular degeneration and is under evaluation for other conditions(27).In the cancer setting, AS1411 targets nucleolin, a protein over-expressed in a variety of tumors. It is currently being evaluated as a potential treatment option in solid tumors and acute myeloid leukemia(28). An updated list of therapeutic aptamers undergoing clinical trials is included in ref.

28 and Table 1. Taken together, these clinical studies highlight many possible uses that aptamers may have in a variety of biomedical fields, including therapeutics.(29)

Table 1

 

A list of therapeutic aptamers undergoing clinical trials

Since aptamer technology was first introduced, the RNA-based sequence library has been widely used for SELEX. Based on the existing evidence, it is believed that the presence of a 2′-OH group and non-Watson-Crick base pairing allows RNA aptamer oligonucleotides to fold into more diverse 3D structures than ssDNA molecules. Consequently, using the more flexible RNA sequences simplifies the development of high-affinity and -specificity aptamers. Despite their advantages, RNA sequences are very sensitive to nucleases present in biological environments and can be rapidly degraded(30). To increase nuclease resistance of RNA-based aptamers, several chemical modifications have been investigated. Evidence shows that 2′-OH group and phosphodiester linkages of RNA sequences are the sites of nuclease hydrolysis. Subsequently, substitutions of the 2′-OH functional group by 2′-fluoro, 2′-amino, or 2′-O-methoxy motifs, and/or changes to the phosphodiester backbone with boranophosphate or phosphorothioate are the most common modifications aimed at increasing nuclease resistance(31). More recently, Wu et al. developed a novel chemical modification method to increase siRNA stability, in which phosphorodithioate and 2′-O-Methyl were simultaneously substituted in the same nucleotide (32). This modification method significantly enhanced siRNA stability and represents a potential new direction for utilization of RNA-based therapies in complex biological systems. Other effective modifications recently reported utilize the locked nucleic acid technology (28, 33) or generate “mirror” RNA sequence structures, termed spiegelmers(34). These modifications result in structural changes to the RNA sequences, which cannot be digested by nucleases.

In addition to RNA aptamers, ssDNA-based aptamers have also been developed. Due to their lack of 2′-OH groups, DNA molecules are naturally resistant to 2′-endonucleases and are stable in biological environments. Recently, our group developed a biostable DNA-based aptamer specific for CD30, a protein biomarker that is over-expressed in Hodgkin and anaplastic large cell lymphomas. Functional analysis demonstrated that this ssDNA-based aptamer exhibited high CD30 binding affinity as low as 2 nmol/l and was stable in human serum for up to 8 hours. Conversely, an RNA-based CD30 aptamer was digested within 10 minutes under similar conditions (35).

In summary, unique chemical features and biological functions have made aptamers a very attractive tool in biomedical research over the past two decades. Currently, there are over 4,000 published articles referenced in the PubMed database that include the term “aptamer.” Research areas that include aptamer technology cover bioassays, drug development, cell detection, tissue staining, in vitro and in vivo imaging, nanotechnology, and targeted therapy. As chemical antibodies, aptamers represent an excellent alternative to replace or supplement protein antibodies, which have been extensively used in the clinic.

 Anti-miRNA Oligonucleotides

Anti-miRNA Oligonucleotides (also known as AMOs) have many uses in cellular mechanics. These synthetically designed molecules are used to neutralize microRNA (miRNA) function in cells for desired responses. miRNA are complementary sequences (≈22 bp) to mRNA that are involved in the cleavage of RNA or the suppression of the translation(36). By controlling the miRNA that regulate mRNAs in cells, AMOs can be used as further regulation as well as for therapeutic treatment for certain cellular disorders. This regulation can occur through a steric blocking mechanism as well as hybridization to miRNA(37). These interactions, within the body between miRNA and AMOs, can be for therapeutics in disorders in which over/under expression occurs or aberrations in miRNA lead to coding issues. Some of the miRNA linked disorders that are encountered in the humans include cancers, muscular diseases, autoimmune disorders, and viruses. In order to determine the functionality of certain AMOs, the AMO/miRNA binding expression (transcript concentration) must be measured against the expressions of the isolated miRNA. The direct detection of differing levels of genetic expression allows the relationship between AMOs and miRNAs to be shown. This can be detected through luciferase activity (biolumincescence in response to targeted enzymatic activity). Understanding the miRNA sequences involved in these diseases can allow us to use anti miRNA Oligonucleotides to disrupt pathways that lead to the under/over expression of proteins of cells that can cause symptoms for these diseases.

 

 RNA interference (RNAi)

RNA interference (RNAi) is a biological process by which double-stranded RNA (dsRNA) inducessequence-specific gene silencing by targeting mRNA for degradation. As a tool for knocking down theexpression of individual genes posttranscriptionally, RNAi has been widely used to study the cellularfunction of genes. In this chapter, I describe procedures for using gene-specific, synthetic, short interferingRNA (siRNA) to induce gene silencing in mammalian cells. Protocols for using lipid-based transfectionreagents and electroporation techniques are provided. Potential challenges and problems associated withthe siRNA technology are also discussed

Specific inhibition or knockdown of gene expression in culturedcells has been widely used to study the effects of loss-of-functionmutation in individual genes. 

 Gene-specific degradation of mRNAis

Gene-specific degradation of mRNAis one way to silence individual gene expression post-transcriptionally. One of the most widely used technologies forinduction of such gene-specific RNA degradation is the use ofRNA interference (RNAi) technology. RNAi was first discoveredin the nematodeC. elegansas a response to small double-strandedRNA (dsRNA), which resulted in sequence-specific genesilencing [38].RNAi is a multistep process. When dsRNA is introduced intocells, it is first recognized and processed into 21–23 base-pair smallinterfering RNAs (siRNA) by Dicer, a RNase III family ribonucle-ase. These short interfering RNAs are then incorporated into anddirect the RNA-induced silencing complex (RISC) to the targetRNA. RISC is a nuclease complex that is responsible for the ulti-mate destruction of the target RNA and gene silencing [39]. In2001, Tuschl and colleagues [40] observed that transfection ofsynthetic 21 base-pair siRNA duplexes into mammalian cells

effectively silences endogenous gene expression in a sequence-specific manner. This finding heralded the use of siRNA for genesilencing in mammalian systems.siRNA oligonucleotides (21–22 base pairs) can be generated bychemical synthesis [41] or by in vitro transcription using T7 RNApolymerase [42]. Alternatively, siRNAs can be endogenouslyexpressed in the form of short hairpin RNA (shRNA), deliveredto cells via plasmids or viral/bacterial vectors [43]. Chemicallysynthesized siRNAs are relatively simple and quick to generate. Inrecent years, a number of commercial manufacturers have started tooffer siRNA oligonucleotide synthesis, which has greatly facilitatedthe use of synthetic siRNAs in research.(44) In this chapter, I will focuson procedures that utilize commercially synthesized siRNAs toknockdown gene expression in mammalian cells.(45)

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