Protein | Turkish Chemistry
May 17

Plasmids

A plasmid is an independent, circular, self-replicating that carries only a few genes. The number of plasmids in a generally remains constant from generation to generation. Plasmids are autonomous and exist in cells as extrachromosomal genomes, although some plasmids can be inserted into a bacterial chromosome, where they become a permanent part of the bacterial genome. It is here that they provide great functionality in molecular science.

Plasmids are easy to manipulate and isolate using bacteria (see also alkaline lysis)

They can be integrated into mammalian genomes, thereby conferring to mammalian cells whatever genetic functionality they carry. Thus, this gives you the ability to introduce genes into a given organism by using bacteria to amplify the hybrid genes that are created in vitro. This tiny but mighty plasmid is the basis of recombinant technology. Plasmids

A plasmid is an independent, circular, self-replicating that carries only a few genes. The number of plasmids in a generally remains constant from generation to generation. Plasmids are autonomous and exist in cells as extrachromosomal genomes, although some plasmids can be inserted into a bacterial chromosome, where they become a permanent part of the bacterial genome. It is here that they provide great functionality in molecular science.

Plasmids are easy to manipulate and isolate using bacteria (see also alkaline lysis) They can be integrated into mammalian genomes, thereby conferring to mammalian cells whatever genetic functionality they carry. Thus, this gives you the ability to introduce genes into a given organism by using bacteria to amplify the hybrid genes that are created in vitro. This tiny but mighty plasmid is the basis of recombinant technology.

There are two categories of plasmids. Stringent plasmids replicate only when the chromosome replicates. This is good if you are working with a that is lethal to the . Relaxed plasmids replicate on their own. This gives you a higher ratio of plasmids to chromosome.

So how do we manipulate these plasmids?

    1. Mutate them using restriction enzymes, ligation enzymes, and PCR. Mutagenesis is easily accomplished by using restriction enzymes to cut out portions of one genome and insert them into a plasmid. PCR can also be used to facilitate mutagenesis. Plasmids are mapped out indicating the locations of their origins of replication and restriction enzyme sites.

    2. Select them using genetic markers. Some bacteria are antibiotic resistant. While this is a serious health problem, it is a godsend to molecular scientists. The gene that confers antibiotic resistance can be added (ligated) to the gene you are inserting into the plasmid. So every plasmid that contains your target gene will not be killed by antibiotics. After you transfect your bacterial cells with your engineered plasmid (the one with the target gene and the antibiotic resistant marker), you incubate them in a nutrient broth that also contains antibiotic (usually ampecillin). Any cells that were not transfected (this means they do not have your target gene in them) are killed by the antibiotic. The ones that do have the gene also have the antibiotic resistant gene, and therefore survive the selection process.

    3. Isolate them (such as with alkaline lysis)

    4. Transform them into cells where they become vectors to transport foreign genes into a recipient organism.

There are some minimum requirements for plasmids that are useful for recombination techniques:

    1. Origin of replication (ORI). They must be able to replicate themselves or they are of no practical use as a vector.

    2. Selectable marker. They must have a marker so you can select for cells that have your plasmids.

    3. Restriction enzyme sites in non-essential regions. You don’t want to be cutting your plasmid in necessary regions such as the ORI.

In addition to these necessary requirements, there are some factors that make plasmids either more useful or easier to work with.

    1. Small. If they are small, they are easier to isolate (you get more), handle (less shearing), and transform.

    2. Multiple restriction enzyme sites. More sites give you greater flexibility in cloning, perhaps even allowing for directional cloning.

    3. Multiple ORIs. It is important to note that two genes must have different ORIs if they are going to be inserted in the same plasmid.

 

May 12

Chromatography

Column chromatography is one of the most common methods of purification. Like many of the techniques on this site, it is as much an art form as a science. Proteins vary hugely in their properties, and the different types of column chromatography allow you to exploit those differences. Most of these methods do not require the denaturing of proteins.

To be very general, a is passed through a column that is designed to trap or slow up the passing of proteins based on a particular property (such as size, charge, or composition).

There are three main steps to purification:

    1. Capture. You need to get your into a concentrated form. If, for example, you are trying to isolate a you have synthesized in an E. coli , you could be looking at a to junk ratio of 1:1,000,000. For capture purification you need a high capacity method that is also fast. You need a speedy method because your crude solution is very likely to contain proteases in addition to your of interest that can quickly chew up your .

    2. Intermediate. Intermediate purification requires both speed and good resolution.

    3. Polishing. For the final step of purification you need a system that has both good resolution and speed. Capacity is usually irrelevant at this stage.

Some of the more common columns include:

  • IEX: Ion exchange chromatography. Good for capture, intermediate, and polish.
  • HIC: Hydrophobic interaction column. Good for intermediate purification.
  • AC: Affinity chromatography. Good for capture and intermediate purification.
  • GF: Gel filtration (size exclusion) chromatography. Good polishing step.

Let’s look at these types of columns in more detail.

Ion exchange chromatography

Ion exchange chromatography is based on the charge of the you are trying to isolate. If your has a high positive charge, you’ll want to pass it through a column with a negative charge. The negative charge on the column will bind the positively charged , and other proteins will pass through the column. You then use a procedure called “salting out” to release your positively charged from the negatively charged column. The column that does this is called a cation exchange column and often uses sulfonated residues. Likewise, you can bind a negatively charged to a positively charge column. The column that does this is called an anion exchange column and often uses quaternary ammonium residues.

Salting out will release, or elute, your from the column. This technique uses a high salt concentration solution. The salt solution will out compete the in binding to the column. In other words, the column has a higher attraction for the charge of salts than for the charged , and it will release the in favor of binding the salts instead. Proteins with weaker ionic interactions will elute at a lower salt, so you will often want to elute with a salt gradient. Different proteins elute at different salt concentrations, so you will want to be sure you know the properties your well for best results.

Also be aware that changes in pH alter the charges in proteins. Be sure you know the isoelectric point of your (the isoelectric point is the pH at which the charge of a is zero) and make sure the pH of your system is adjusted and buffered accordingly.

The basic steps in using an ion exchange column are:

    1. Prep the column. Pour your buffer over the column to make sure it has equilibrated to the required pH.

    2. Load your solution. Some proteins in the solution don’t bind and will elute during this loading phase.

    3. Salt out. Increase the salt concentration to elute the bound proteins. It is best to use a salt gradient to gradually elute proteins with different ionic strengths. At the end bump the system with a very high salt concentration (2-3M) to make sure all proteins are off the column.

    4. Remove salts. Use dialysis to remove the salts from your solution.

Temperature doesn’t have a huge effect on column chemistry. However, it is better to work cold since proteins are more stable cold.

Hydrophobic interaction chromatography

Where ion exchange chromatography relies on the charges of proteins to isolate them, hydrophobic interaction chromatography uses the hydrophobic properties of some proteins. Hydrophobic groups on the bind to hydrophillic groups on the column. The more hydrophobic a is, the stronger it will bind to the column.

Load the proteins in the presence of a high concentration of ammonium sulfate (not ammonium persulfate). Ammonium sulfate is a chaotropic agent. It increases the chaos (entropy) in , and thereby increases hydrophobic interactions (the more disordered the , the stronger the hydrophobic interactions). Ammonium sulfate also stabilizes proteins. So as a result of using an HIC column you can expect your to be in its most stable form.

The hydrophobic column is packed with a phenyl agarose matrix. In the presence of high salt concentrations the phenyl groups on this matrix binds hydrophobic portions of proteins. You can control elution of different column-bound proteins by reducing the salt concentration or by adding solvents.

Affinity chromatography.

Affinity chromatography relies on the biological functions of a to bind it to a column. The most common type involves a ligand, a specific small biomolecule. This small molecule is immobilized and attached to a column matrix, such as cellulose or polyacrylamide. Your target is then passed through the column and bound to it by its ligand, while other proteins elute out. Elution of your target is usually done by passing through the column a solution that has in it a high concentration of free ligand.  This is a very efficient purification method since it relies on the biological specificity of your target , such as the affinity of an enzyme for a substrate.

Gel filtration, or size exclusion, chromatography separates proteins on the basis of their size. The column is packed with a matrix of fine porous beads.

It works somewhat like a sieve, but in reverse. The beads have in them very small holes. As the solution is poured on the column, small enter the pores in the beads. Larger are excluded from the holes, and pass quickly between the beads.

These larger are eluted first. The smaller have a longer path to travel, as they get stuck over and over again in the maze of pores running from bead to bead. These smaller , therefore, take longer to make their way through the column and are eluted last.

May 11

Gel electrophoresis

Gel electrophoresis is a technique used for the separation of deoxyribonucleic (DNA), ribonucleic (RNA), or protein using an electric current applied to a gel matrix.[1] It is usually performed for analytical purposes, but may be used as a preparative technique prior to use of other methods such as mass spectrometry, RFLP, , cloning, DNA sequencing, or Southern blotting for further characterization.

The term “gel” in this instance refers to the matrix used to contain, then separate the target . In most cases, the gel is a crosslinked polymer whose composition and porosity is chosen based on the specific weight and composition of the target to be analyzed. When separating proteins or small nucleic acids (DNA, RNA, or oligonucleotides) the gel is usually composed of different concentrations of acrylamide and a cross-linker, producing different sized mesh networks of polyacrylamide. When separating larger nucleic acids (greater than a few hundred bases), the preferred matrix is purified agarose. In both cases, the gel forms a solid, yet porous matrix. Acrylamide, in contrast to polyacrylamide, is a neurotoxin and must be handled using appropriate safety precautions to avoid poisoning. Another gel matrix is agarose, long unbranched chains of uncharged carbohydrate without cross links giving a gel with large pores allowing separation of macromolecules and macromolecular complexes.

“Electrophoresis” refers to the electromotive force (EMF) that is used to move the through the gel matrix. By placing the in wells in the gel and applying an electric current, the will move through the matrix at different rates, determined largely by their mass when the charge to mass ratio (Z) of all species is uniform, toward the anode if negatively charged or toward the cathode if positively charged

After the electrophoresis is complete, the in the gel can be stained to make them visible. Ethidium bromide, silver, or coomassie blue dye may be used for this process. Other methods may also be used to visualize the separation of the mixture’s components on the gel. If the analyte fluoresce under ultraviolet light, a photograph can be taken of the gel under ultraviolet lighting conditions. If the to be separated contain radioactivity added for visibility, an autoradiogram can be recorded of the gel.

If several mixtures have initially been injected next to each other, they will run parallel in individual lanes. Depending on the number of different , each lane shows separation of the components from the original mixture as one or more distinct bands, one band per component. Incomplete separation of the components can lead to overlapping bands, or to indistinguishable smears representing multiple unresolved components.

Bands in different lanes that end up at the same distance from the top contain that passed through the gel with the same speed, which usually means they are approximately the same size. There are molecular weight size markers available that contain a mixture of of known sizes. If such a marker was run on one lane in the gel parallel to the unknown samples, the bands observed can be compared to those of the unknown in order to determine their size. The distance a band travels is approximately inversely proportional to the logarithm of the size of the molecule.

Applications

Gel electrophoresis is used in forensics, molecular biology, genetics, microbiology and . The results can be analyzed quantitatively by visualizing the gel with UV light and a gel imaging device. The image is recorded with a computer operated camera, and the intensity of the band or spot of interest is measured and compared against standard or markers loaded on the same gel. The measurement and analysis are mostly done with specialized software.

Depending on the type of analysis being performed, other techniques are often implemented in conjunction with the results of gel electrophoresis, providing a wide range of field-specific applications.

Nucleic acids

In the case of nucleic acids, the direction of migration, from negative to positive electrodes, is due to the naturally-occurring negative charge carried by their sugar-phosphate backbone.[3]

Double-stranded DNA fragments naturally behave as long rods, so their migration through the gel is relative to their radius of gyration, or, for non-cyclic fragments, size. Single-stranded DNA or RNA tend to fold up into with complex shapes and migrate through the gel in a complicated manner based on their tertiary structure. Therefore, agents that disrupt the hydrogen bonds, such as sodium hydroxide or formamide, are used to denature the nucleic acids and cause them to behave as long rods again.[4]

Gel electrophoresis of large DNA or RNA is usually done by agarose gel electrophoresis. See the “Chain termination method” page for an example of a polyacrylamide DNA sequencing gel. Characterization through ligand interaction of nucleic acids or fragments may be performed by mobility shift affinity electrophoresis.

Proteins

SDS-PAGE autoradiography – The indicated proteins are present in different concentrations in the two samples.

Proteins, unlike nucleic acids, can have varying charges and complex shapes, therefore they may not migrate into the polyacryl amide gel at similar rates, or at all, when placing a negative to positive EMF on the sample. Proteins therefore, are usually denatured in the presence of a detergent such as sodium dodecyl sulfate/sodium dodecyl phosphate (SDS/SDP) that coats the proteins with a negative charge.[1] Generally, the amount of SDS bound is relative to the size of the protein (usually 1.4g SDS per gram of protein), so that the resulting denatured proteins have an overall negative charge, and all the proteins have a similar charge to mass ratio. Since denatured proteins act like long rods instead of having a complex tertiary shape, the rate at which the resulting SDS coated proteins migrate in the gel is relative only to its size and not its charge or shape.[1]

Proteins are usually analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), by native gel electrophoresis, by quantitative preparative native continuous polyacrylamide gel electrophoresis (QPNC-PAGE), or by 2-D electrophoresis.

Characterization through ligand interaction may be performed by electroblotting or by affinity electrophoresis in agarose or by capillary electrophoresis as for estimation of binding constants and determination of structural features like glycan content through lectin binding.

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