Molecular Techniques

Inactivating and Marking Target Genes with Transposons

Transposons allow genes to be transferred to a host organism’s chromosome, interrupting or modifying the function of a gene.

Learning Objectives

Describe the utility of experimentally introduced Transposons

Key Takeaways

Key Points

  • Transposons contain signals to truncate expression of an interrupted gene, thus inactivating it.
  • Transposons are widely used tools in biology, frequently utilized for insertion mutagenesis, large-scale gene disruption studies, and gene tagging.
  • Transposon-mediated gene disruption experiments and promoter traps rely on promiscuous, undirected, and pseudo-random insertion of the transposon.

Key Terms

  • transposable: Able to be transposed (in any sense).
  • plasmid: A circle of double-stranded DNA that is separate from the chromosomes, which is found in bacteria and protozoa.

A transposable element (TE) is a DNA sequence that can change its relative position (self-transpose) within the genome of a single cell. The mechanism of transposition can be either “copy and paste” or “cut and paste. ” Transposition can create phenotypically significant mutations and alter the cell’s genome size. Barbara McClintock’s discovery of these jumping genes early in her career earned her a Nobel prize in 1983.

Transposons in bacteria usually carry an additional gene for function other than transposition—often for antibiotic resistance. In bacteria, transposons can jump from chromosomal DNA to plasmid DNA and back, allowing for the transfer and permanent addition of genes such as those encoding antibiotic resistance (multi-antibiotic resistant bacterial strains can be generated in this way). When the transposable elements lack additional genes, they are known as insertion sequences. Transposons are semi-parasitic DNA sequences that can replicate and spread through the host ‘s genome. They can be harnessed as a genetic tool for analysis of gene and protein function. The use of transposons is well-developed in Drosophila (in which P elements are most commonly used) and in Thale cress (Arabidopsis thaliana) and bacteria such as Escherichia coli (E. coli ).

Synthetic DNA transposon system are constructed to introduce precisely defined DNA sequences into the chromosomes of vertebrate animals for the purposes of introducing new traits and to discover new genes and their functions (e.g. by establishing a loss-of-function phenotype or gene inactivation). Transposition is a precise process in which a defined DNA segment is excised from one DNA molecule and moved to another site in the same or different DNA molecule or genome.

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Transposon System: The sleeping beauty transposon system applications.

Insertional inactivation is a technique used in recombinant DNA engineering where a plasmid (such as pBR322) is used to disable the expression of a gene. A gene is inactivated by inserting a fragment of DNA into the middle of its coding sequence. Any future products from the inactivated gene will not work because of the extra codes added to it. An example is the use of pBR322, which has genes that respectively encode polypeptides that confer resistance to ampicillin and tetracyclin antibiotics. As a result, when a genetic region is interrupted by integration of pBR322, the gene function is lost but new gene function (resistance to specific antibiotics) is gained. An alternative strategy for insertional mutagenesis has been used in vertebrate animals to find genes that cause cancer. In this case a transposon, e.g. Sleeping Beauty, is designed to interrupt a gene in such a way that it causes maximal genetic havoc. Specifically, the transposon contains signals to truncate expression of an interrupted gene at the site of the insertion and then restart expression of a second truncated gene. This method has been used to identify oncogenes.

DNA Sequencing of Insertion Sites

An insertion site is the position at which a transposable genetic element is integrated.

Learning Objectives

Discuss the uses of sequencing insertions sites

Key Takeaways

Key Points

  • Several methods exist to analyze insertion sequences, including inverse polymerase chain reaction. The inverse PCR involves a series of restriction digests and ligation, resulting in a looped fragment that can be primed for PCR from a single section of known sequence.
  • The amplified product can then be sequenced and compared with DNA databases to locate the sequence which has been disrupted.
  • Other techniques include Southern hybridization and modifications of the PCR protocol.

Key Terms

  • transposons: A segment of DNA that can move to a different position within a genome.

An insertion sequence (also known as an IS, an insertion sequence element, or an IS element) is a short DNA sequence that acts as a simple transposable element.

Insertion sequences have two major characteristics: they are small relative to other transposable elements (generally around 700 to 2500 bp in length) and only code for proteins implicated in the transposition activity (they are thus different from other transposons, which also carry accessory genes such as antibiotic-resistance genes).

These proteins are usually the transposase which catalyse the enzymatic reaction allowing the IS to move, and also one regulatory protein which either stimulates or inhibits the transposition activity. The coding region in an insertion sequence is usually flanked by inverted repeats. For example, the well-known IS911 (1250 bp) is flanked by two 36bp inverted repeat extremities and the coding region has two genes partially overlapping orfA and orfAB, coding the transposase (OrfAB) and a regulatory protein (OrfA).

A particular insertion sequence may be named according to the form ISn, where n is a number (e.g. IS1, IS2, IS3, IS10, IS50, IS911, IS26, etc.); this is not the only naming scheme used, however. Although insertion sequences are usually discussed in the context of prokaryotic genomes, certain eukaryotic DNA sequences belonging to the family of Tc1/mariner transposable elements may be considered to be insertion sequences.

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Naegleria fowleri: Antibody detection (green) of Naegleria fowleri, the organism responsible for Primary amoebic meningoencephalitis (PAM).

In addition to occurring autonomously, insertion sequences may also occur as parts of composite transposons; in a composite transposon, two insertion sequences flank one or more accessory genes, such as an antibiotic-resistance gene (e.g. Tn10, Tn5). Nevertheless, there exist another sort of transposons, called unit transposons, that do not carry insertion sequences at their extremities (e.g. Tn7). A complex transposon does not rely on flanking insertion sequences for resolvase. The resolvase is part of the tns genome and cuts at flanking inverted repeats.

Although several methods are available for locating ISs in microbial genomes, they are either labor intensive or inefficient. These include Southern hybridization, inverse Polymerase Chain Reaction (iPCR), and most recently, vectorette PCR to identify and map the genomic positions of the insertion sequences.

Southern hybridization is rather time-consuming and requires additional procedures for localizing ISs. Inverse PCR, a commonly-used PCR method for recovering unknown flanking sequences of a known target sequence, uses a library of circularized chromosomal DNA fragments as a template and two outward primers located in each end of the known fragment for amplification. However, when a target sequence has multiple genomic locations, the variously-sized DNA circles formed are difficult to amplify simultaneously. Also, the length of each restriction DNA fragment containing a target sequence must be determined by Southern hybridization followed by sub-genomic fractioning before intramolecular ligation and PCR amplification. These difficulties render Southern hybridization and iPCR impractical as techniques for quickly surveying repetitive elements in genomes.

Vectorette PCR (vPCR) is another method used to amplify unknown sequences flanking a characterized DNA fragment. It involves cutting genomic DNAs with a restriction enzyme, ligating vectorettes to the ends, and amplifying the flanking sequences of a known sequence using primers derived from the known sequence along with a vectorette primer.

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Inverse PCR: Summary of iPCR process

Northern Blots

Northern blots allow investigators to determine messenger RNA molecular weight and sample content.

Learning Objectives

Evaluate the applications of Northern Blots

Key Takeaways

Key Points

  • RNA (either total RNA or just mRNA) is separated by gel electrophoresis, usually an agarose gel. Because there are so many different RNA molecules on the gel, it usually appears as a smear rather than discrete bands.
  • The RNA is transfered to a sheet of special blotting paper called nitrocellulose, though other types of paper, or membranes, can be used. The RNA molecules retain the same pattern of separation they had on the gel.
  • The blot is incubated with a probe which is single-stranded DNA. This probe will form base pairs with its complementary RNA sequence and bind to form a double-stranded RNA-DNA molecule. The probe is either radioactive or has an enzyme bound to it.

Key Terms

  • hybridization: The act of hybridizing, or the state of being hybridized.

The Northern blot is a technique used in molecular biology research to study gene expression in a sample, through detection of RNA (or isolated messenger RNA ). With Northern blotting it is possible to observe cellular control over structure and function by determining the particular gene expression levels during differentiation, morphogenesis, as well as abnormal or diseased conditions. Northern blotting involves the use of electrophoresis to separate RNA samples by size and detection with a hybridization probe complementary to part of or the entire target sequence.

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Northern blot technique: Flow diagram outlining the general procedure for RNA detection by northern blotting.

The term ‘Northern blot’ actually refers specifically to the capillary transfer of RNA from the electrophoresis gel to the blotting membrane. However, the entire process is commonly referred to as Northern blotting. The northern blot technique was developed in 1977 by James Alwine, David Kemp, and George Stark at Stanford University. Northern blotting takes its name from its similarity to the first blotting technique, the Southern blot, named for biologist Edwin Southern. The major difference is that RNA, rather than DNA, is analyzed in the Northern blot.

A general blotting procedure starts with extraction of total RNA from a homogenized tissue sample or from cells. Eukaryotic mRNA can then be isolated through the use of oligo (dT) cellulose chromatography to isolate only those RNAs with a poly(A) tail. RNA samples are then separated by gel electrophoresis. Since the gels are fragile and the probes are unable to enter the matrix, the RNA samples, now separated by size, are transferred to a nylon membrane through a capillary or vacuum blotting system.

Western Blots

The Western blot technique determines protein molecular weight and measures protein abundance in different samples.

Learning Objectives

Show the uses of Western Blots

Key Takeaways

Key Points

  • After separation by gel electrophoresis using SDS-PAGE, proteins are transfered to a sheet of special blotting paper called nitrocellulose, though other types of paper, or membranes, can be used. The proteins retain the same pattern of separation they had on the gel.
  • The blot is incubated with a generic protein (such as milk proteins) to bind to any remaining sticky places on the nitrocellulose. An antibody is then added to the solution which is able to bind to its specific protein. The antibody is conjugated to alkaline phosphatase or horseradish peroxidase.
  • The location of the antibody is revealed by incubating it with a colorless substrate that the attached enzyme converts to a colored product that can be seen and photographed.

Key Terms

  • electrophoresis: a method for the separation and analysis of large molecules (such as proteins) by migrating a colloidal solution of them through a gel; gel electrophoresis

The Western blot (sometimes called the protein immunoblot) is a widely accepted analytical technique used to detect specific proteins in a given sample of tissue homogenate or extract. Western blot samples can be taken from whole tissue or from cell culture. Solid tissues are first broken down mechanically using either a blender (for larger sample volumes), a homogenizer (smaller volumes), or by sonication. Assorted detergents, salts, and buffers may be employed to encourage lysis of cells and to solubilize proteins. The technique uses gel electrophoresis to separate native proteins by 3-D structure or denatured proteins by the length of the polypeptide.

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Western blot steps: Example preparation to use in the Western blot technique.

The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are stained with antibodies specific to the target protein. There are now many reagent companies that specialize in providing antibodies (both monoclonal and polyclonal antibodies) against tens of thousands of different proteins belonging to signaling pathways or cell surface receptor antigens, or other cellular or soluble components. Commercial antibodies can be expensive, although the unbound antibody can be reused between experiments. This method is used in the fields of molecular biology, biochemistry, immunogenetics and other molecular biology disciplines. Other related techniques include using antibodies to detect proteins in tissues and cells by immunostaining and enzyme-linked immunosorbent assay (ELISA). This method originated in the laboratory of George Stark at Stanford. The name Western blot was given to the technique by W. Neal Burnette and is a play on the name Southern blot, a technique for DNA detection developed earlier by Edwin Southern. Detection of RNA is termed Northern blot.

DNA Mobility Shifts

DNA mobility shift assay is a technique for studying gene regulation and determining protein-DNA interactions.

Learning Objectives

Identify the utility of DNA mobility shift assays

Key Takeaways

Key Points

  • The interaction of proteins with DNA is central to the control of many cellular processes, including DNA replication, recombination and repair, transcription, and viral assembly.
  • An advantage of studying protein-DNA interactions by an electrophoretic mobility shift assay is the ability to resolve complexes of different stoichiometry or conformation.
  • The source of the DNA-binding protein may be a crude nuclear or whole cell extract, in vitro transcription product, or a purified preparation.

Key Terms

  • polyacrylamide: Any of a range of cross-linked polymers of acrylamide; used to form soft gels.

A mobility shift assay is electrophoretic separation of a protein-DNA or protein- RNA mixture on a polyacrylamide or agarose gel for a short period. The speed at which different molecules (and combinations thereof) move through the gel is determined by their size and charge, and to a lesser extent, their shape. The control lane (a DNA probe without protein present) will contain a single band corresponding to the unbound DNA or RNA fragment. However, assuming that the protein is capable of binding to the fragment, the lane with protein present will contain another band that represents the larger, less mobile, complex of nucleic acid probe bound to protein, which is “shifted” up on the gel (since it has moved more slowly).

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Gel Shift Assay: Lane 1 is a negative control, and contains only DNA. Lane 2 contains protein as well as a DNA fragment that, based on its sequence, does not interact. Lane 3 contains protein and a DNA fragment that does react; the resulting complex is larger, heavier, and slower-moving. The pattern shown in lane 3 is the one that would result if all the DNA were bound and no dissociation of complex occurred during electrophoresis. When these conditions are not met a second band might be seen in lane 3 reflecting the presence of free DNA or the dissociation of the DNA-protein complex.

Under the correct experimental conditions, the interaction between the DNA and protein is stabilized and the ratio of bound to unbound nucleic acid on the gel reflects the fraction of free and bound probe molecules as the binding reaction enters the gel. This stability is in part due to the low ionic strength of the buffer, but also due to a “caging effect”; the protein, surrounded by the gel matrix, is unable to diffuse away from the probe before they recombine. If the starting concentrations of protein and probe are known, and if the stoichiometry of the complex is known, the apparent affinity of the protein for the nucleic acid sequence may be determined. An antibody that recognizes the protein can be added to this mixture to create an even larger complex with a greater shift. This method is referred to as a supershift assay, and is used to unambiguously identify a protein present in the protein-nucleic acid complex.

Purifying Proteins by Affinity Tag

Protein tags are peptide sequences genetically grafted onto a recombinant protein.

Learning Objectives

Indicate the uses of protein affinity tags

Key Takeaways

Key Points

  • Affinity tags are appended to proteins so that they can be purified from their crude biological source using an affinity technique.
  • Recombinant proteins that carry small affinity tags are efficiently expressed in bacteria, insect, or mammalian cells.
  • After cell lysis and clearing of the lysate, tagged proteins are purified using an immobilized-metal affinity chromatography procedure.

Key Terms

  • protein: Proteins are large biological molecules consisting of one or more chains of amino acids.
  • affinity: An attractive force between atoms, or groups of atoms, that contributes toward their forming bonds.
  • recombinant: This term refers to something formed by combining existing elements in a new combination. Thus, the phrase recombinant DNA refers to an organism created in the lab by adding DNA from another species.

Protein tags are peptide sequences genetically grafted onto a recombinant protein. Often these tags are removable by chemical agents or by enzymatic means, such as proteolysis or intein splicing. Tags are attached to proteins for various purposes.

Affinity tags are appended to proteins so that they can be purified from their crude biological source using an affinity technique. These include chitin binding protein (CBP), maltose binding protein (MBP), and glutathione-S-transferase (GST). The poly (His) tag is a widely-used protein tag; it binds to metal matrices.

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Adding Polyhistidine Tags: This is an example of a primer designed to add a 6xHis-tag using PCR. Eighteen bases coding six histidines are inserted right after the START codon or right before the STOP codon. At least 16 bases specific to the gene of interest are needed next to the His-tag. With 6 His, the protein will have an added 1 kDa of molecular weight. Oftentimes, a linker (such as gly-gly-gly or gly-ser-gly) is placed between the protein of interest and the 6 His tag. This is to prevent the polyhistidine tag from affecting the activity of the protein being tagged.

Solubilization tags are used, especially for recombinant proteins expressed in chaperone-deficient species such as E. coli, so as to assist in the proper folding in proteins and keep them from precipitating. These include thioredoxin (TRX) and poly (NANP). Some affinity tags have a dual role as a solubilization agent, such as MBP and GST.

Chromatography tags are used to alter chromatographic properties of the protein to afford different resolution across a particular separation technique. These often consist of polyanionic amino acids, such as FLAG-tag.

Epitope tags are short peptide sequences which are chosen because high-affinity antibodies can be reliably produced in many different species. These are usually derived from viral genes, which explain their high immunoreactivity. Epitope tags include V5-tag, c-myc-tag, and HA-tag. These tags are particularly useful for western blotting, immunofluorescence and immunoprecipitation experiments, although they also find use in antibody purification.

Fluorescence tags are used to give visual readout on a protein. GFP and its variants are the most commonly used fluorescence tags. More advanced applications of GFP include using it as a folding reporter (fluorescent if folded, colorless if not).

Protein tags are also useful for specific enzymatic modification (such as biotin ligase tags) and chemical modification (FlAsH) tag. Often tags are combined to produce multifunctional modifications of the protein. However, with the addition of each tag comes the risk that the native function of the protein may be abolished or compromised by interactions with the tag.

Examples of peptide tags include:

  • AviTag, a peptide allowing biotinylation by the enzyme BirA and so the protein can be isolated by streptavidin (GLNDIFEAQKIEWHE)
  • Calmodulin-tag, a peptide bound by the protein calmodulin (KRRWKKNFIAVSAANRFKKISSSGAL)
  • FLAG-tag, a peptide recognized by an antibody (DYKDDDDK)
  • HA-tag, a peptide recognized by an antibody (YPYDVPDYA)
  • His-tag, 5-10 histidines bound by a nickel or cobalt chelate (HHHHHH)
  • Myc-tag, a short peptide recognized by an antibody (EQKLISEEDL)
  • S-tag (KETAAAKFERQHMDS)
  • SBP-tag, a peptide which binds to streptavidin (MDEKTTGWRGGHVVEGLAGELEQLRARLEHHPQGQREP)
  • Softag 1, for mammalian expression (SLAELLNAGLGGS)
  • Softag 3, for prokaryotic expression (TQDPSRVG)
  • V5 tag, a peptide recognized by an antibody (GKPIPNPLLGLDST)
  • Xpress tag (DLYDDDDK)

Examples of protein tags include:

  • BCCP (Biotin Carboxyl Carrier Protein), a protein domain recognized by streptavidin
  • Glutathione-S-transferase-tag, a protein which binds to immobilized glutathione
  • Green fluorescent protein-tag, a protein which is spontaneously fluorescent and can be bound by nanobodies
  • Maltose binding protein-tag, a protein which binds to amylose agarose
  • Nus-tag
  • Strep-tag, a peptide which binds to streptavidin or the modified streptavidin called streptactin (Strep-tag II: WSHPQFEK)
  • Thioredoxin-tag

Primer Extension Analysis

Primer extension is used to map the 5′ ends of DNA or RNA fragments.

Learning Objectives

Outline primer extension analysis

Key Takeaways

Key Points

  • Primer extension assay is done by annealing a specific oligonucleotide primer to a position downstream of that 5′ end.
  • The primer is radiolabelled, usually at its 5′ end. This is extended with reverse transcriptase, which can copy either an RNA or a DNA template, making a fragment that ends at the 5′ end of the template molecule.
  • Primer extension analysis includes selection and preparation of a labeled primer complementary to the RNA transcript of interest; hybridization of the primer to a region of the RNA under study; extension from the primer by an RNA-dependent DNA polymerase to synthesize a cDNA strand.
  • Analysis of primer extension of the extended cDNA products is done on denaturing polyacrylaminde gels and autoradiography.

Key Terms

  • radiolabelled: Tagged with a radiotracer.
  • polyacrylamide: Any of a range of cross-linked polymers of acrylamide; used to form soft gels.

Primer Extension Analysis

Primer extension is a technique whereby the 5′ ends of RNA or DNA can be mapped. Primer extension can be used to determine the start site of RNA transcription for a known gene. This technique requires a radiolabelled primer (usually 20 to 50 nucleotides in length) which is complementary to a region near the 3′ end of the gene. The primer is allowed to anneal to the RNA and reverse transcriptase is used to synthesize cDNA from the RNA until it reaches the 5′ end of the RNA. By running the product on a polyacrylamide gel, it is possible to determine the transcriptional start site, as the length of the sequence on the gel represents the distance from the start site to the radiolabelled primer.

Applications of Primer Extension Analysis

Primer extension analysis has three main applications. First, it is used for mapping the 5′ end of transcripts. This allows one to determine the transcription initiation site (assuming the mRNA isn’t further processed), which helps localize promoters or TATA boxes. Second, it can be used to quantify the amount of transcript in an in vitro transcription system.

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Transcription initiation site: This is a diagram of transcription initiation by the RNA polymerase.

Third, it can be used to determine the locations of breaks or modified bases in a mixed population of RNA or DNA samples. This is useful in applications like footprinting. Two different methods are used. In one, the modified nucleotide cannot be recognized by the polymerase or reverse transcriptase; in such cases, the chain ends at the site of modification. In the other, the modification is converted in a later step of the analysis to a strand break by chemical treatment. For instance, the sites of modifications by dimethyl sulfate (DMS) can be identified by treating DNA with DMS, exposing the sample to conditions that break the backbone at the site of modification, followed by primer extension.

DNA Protection Analysis

DNA protection or “footprinting” analysis is a powerful technique for identifying the nucleotides involved in a protein-DNA interaction.

Learning Objectives

Illustrate DNA protection analysis

Key Takeaways

Key Points

  • DNA protection analysis is a technique in which a DNA molecule is ‘incubated’ with a protein that binds to a specific site along the double helix.
  • The DNA-binding protein complex is then subjected to restriction endonuclease digestion, which reduces the entire DNA to mono- and oligonucleotide fragments, except for the portion of the DNA molecule that was ‘protected’ from digestion by the binding protein.
  • Removal of the protein by simple chemical means—e.g., by gel electrophoresis —allows the study of DNA and binding protein interaction.

Key Terms

  • electrophoresis: a method for the separation and analysis of large molecules (such as proteins) by migrating a colloidal solution of them through a gel; gel electrophoresis
  • polymerase chain reaction: A technique in molecular biology for creating multiple copies of DNA from a sample; used in genetic fingerprinting etc.

DNA protection or footprinting is a technique from molecular biology/biochemistry that detects DNA-protein interaction using the fact that a protein bound to DNA will often protect that DNA from enzymatic cleavage. This makes it possible to locate a protein binding site on a particular DNA molecule. The method uses an enzyme, deoxyribonuclease (DNase, for short) to cut the radioactively end-labeled DNA, followed by gel electrophoresis to detect the resulting cleavage pattern. For example, the DNA fragment of interest may be amplified by polymerase chain reaction, with the result being many DNA molecules with a radioactive label on one end of one strand of each double stranded molecule. Cleavage by DNase will produce fragments, the smaller of which will move further on the electrophoretic gel.

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DNA footprinting: DNA protection or footprinting technique

The fragments which are smaller will appear further on the gel than the longer fragments. The gel is then used to expose a special photographic film. The cleavage pattern of the DNA in the absence of a DNA binding protein, typically referred to as free DNA, is compared to the cleavage pattern of DNA in the presence of a DNA binding protein. If the protein binds DNA, the binding site is protected from enzymatic cleavage. This protection will result in a clear area on the gel which is referred to as the “footprint”. By varying the concentration of the DNA-binding protein, the binding affinity of the protein can be estimated according to the minimum concentration of protein at which a footprint is observed. This technique was developed by David Galas and Albert Schmitz at Geneva in 1977.

Whole-Genome DNA-Binding Analysis

Whole-genome DNA-binding analysis is a powerful tool for analyzing epigenetic modifications and DNA sequences bound to regulatory proteins.

Learning Objectives

Describe whole-genome DNA-binding analysis

Key Takeaways

Key Points

  • Whole-genome DNA binding analysis, also known as location analysis, utilizes chromatin immunoprecipitation and microarray chip.
  • Whole-genome DNA-binding analysis utilizes ChIP-on-chip method. Briefly, protein -DNA complexes are crosslinked, immunoprecipitated, purified, amplified and labeled, and then allowed to hybridize to a variety of high-resolution arrays.
  • This technique is a high-throughput (genome-wide) identification and analysis of DNA fragments that are bound by specific proteins such as histones, and transcriptional factors.

Key Terms

  • immunoprecipitation: A technique in which an antigen is precipitated from a solution by using an antibody, or a particular use of this technique.

Genomic DNA sequences are being determined at an increasingly rapid pace. This has created a need for more efficient techniques to determine which parts of these sequences are bound in-vivo by the proteins controlling processes; such as gene expression, DNA replication and chromosomal mechanics.

A whole-genome approach was established to identify and characterize such DNA sequences. The method of chromatin immunoprecipitation, combined with microarrays (ChIP-Chip), is a powerful tool for genome-wide analysis of protein binding. It has also become a widely-used method for genome-wide localization of protein-DNA interactions.

The first step in the ChIP-Chip procedure is to fix protein-DNA interactions in living cells by chemical crosslinking. The crosslinker must be small to diffuse fast into the cells. In practice, formaldehyde is used in most ChIP-Chip experiments. After cell lysis, the DNA is fragmented by sonication. This extract is then subjected to immunoprecipitation (IP) with a specific antibody against the protein of interest.

DNA bound by the protein will be coprecipitated and enriched, compared to DNA not bound by the respective protein. To facilitate immunoprecipitation and subsequent washing, antibodies are usually coupled to either agarose- or magnetic beads via protein A or G. After reversion of crosslinking, the DNA is purified by phenol extraction or commercial polymerase chain reaction (PCR) cleanup kits.

Often, an amplification step is included after DNA purification. Two different fluorescence labels are used to label the IP DNA, and a hybridization -control DNA, respectively. Usually, total DNA before IP (input DNA) is used as hybridization control.

The two differentially-labeled DNAs are hybridized to the same microarray and the difference in fluorescence intensity gives a measure of the enrichment.

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ChIP-chip procedure: ChIP-chip workflow performed in laboratory

Two-Hybrid Analysis

The two-hybrid method detects the interaction of two proteins by their ability to reconstitute the activity of a split transcription factor.

Learning Objectives

Design a two-hybrid experiment

Key Takeaways

Key Points

  • A key part of gene functional analysis and potential drug target discovery is an understanding of how proteins interact within the cell. Commercially available products facilitate the characterization of these interactions in yeast systems.
  • The basic format involves the creation of two hybrid molecules, one in which a “bait” protein is fused with a transcription factor, and one in which a “prey” protein is fused with a related transcription factor.
  • If the bait and prey proteins indeed interact, then the two factors fused to these two proteins are also brought into proximity with each other.

Key Terms

  • computational: Of or relating to computation.
  • ubiquitin: A small regulatory protein sequence that directs proteins to specific compartments within the cell. Specifically, a ubiquitin tag directs the protein to a proteasome, which destroys and recycles the components.

Understanding how proteins are physically connected reveals clues about their structure, function, and makes them an ideal target for drug therapy. Several methodologies exist to study the interaction of proteins in vivo. The most widely employed tools are the yeast two-hybrid system. The yeast two-hybrid screening system is an effective and quick tool for the in vivo study of protein–protein interaction both in prokaryotes and eukaryotes. The method consists of splitting a yeast transcription factor into its binding domain and activation domain, fusing the binding domain to one protein of interest (the bait) and the activation domain to another protein of interest (the prey), and reconstituting the activity of the transcription factor by bringing the two domains back into physical proximity. In the absence of an interaction the domains remain distant, preventing a detectable output. If the two proteins do interact the bait recruits the prey to a specific cellular location where it can stimulate a detectable output (e.g., gene activation). This experimental approach measures direct physical interaction between proteins and is called a binary method. Datasets obtained from such tools are further analyzed using computational methods to draw a map of protein connectivity and achieve system level understanding of a microorganism.

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Two-hybrid technique: Overview of two-hybrid assay, checking for interactions between two proteins, called here Bait and Prey.

One limitation of classic yeast two-hybrid screens is that they are limited to soluble proteins. It is therefore impossible to use them to study the protein–protein interactions between insoluble integral membrane proteins. The split- ubiquitin system provides a method for overcoming this limitation. In the split-ubiquitin system, two integral membrane proteins to be studied are fused to two different ubiquitin moieties: a C-terminal ubiquitin moiety (“Cub”, residues 35–76) and an N-terminal ubiquitin moiety (“Nub”, residues 1–34). These fused proteins are called the bait and prey, respectively. In addition to being fused to an integral membrane protein, the Cub moiety is also fused to a transcription factor (TF) that can be cleaved off by ubiquitin specific proteases. Upon bait–prey interaction, Nub and Cub-moieties assemble, reconstituting the split-ubiquitin. The reconstituted split-ubiquitin molecule is recognized by ubiquitin specific proteases, which cleave off the reporter protein, allowing it to induce the transcription of reporter genes.