Laboratory Practices - GRE Subject Test: Biochemistry, Cell, and Molecular Biology

FRET is a widely used technique to study protein-protein interactions as well as reaction kinetics. The technique works by attaching fluorophores to proteins that produce fluorescent light when stimulated by light of the appropriate wavelength. When attempting to detect distances between two objects, two fluorophores are used. The first one is stimulated by an external source and the second is stimulated by the light produced from the excited fluorophore. The presence or absence of the light produced by the second excited fluorophore can then help determine protein-protein interactions. The technique is also applicable to protein localization and detecting distances between domains.

During FRAP experimentation, a section of membrane is labeled with fluorophores. High intensity exposure to a small region of the membrane is used to "bleach" this region, resulting in a small region of zero fluorescence. The lack of recovery of the fluorescence only gives us information about the mobility of the protein: it is immobile. If the protein were mobile, then tagged proteins would be able to move into the bleached region, returning it to fluorescence.

While it may be true that the protein is anchored to the cytoskeleton or part of a lipid raft, the result of the experiment does not directly lead to these conclusions. It does, however, point the researcher in the right direction about learning more about the functionality of his protein. FRAP does not tell us anything about the stability of the protein.

GFP is a fluorescent tag that is incredibly useful in helping visualize a protein in vivo. The tag will glow green in the tissue where the protein is present.

All of the other choices describe tags that help purify recombinant proteins. Because all of these tags isolate proteins from a sample of lysed cell, they would not be useful in following a protein in vivo.

EDTA is a chelator of metal cations. In order to detect a 6-His tag, nickel beads are used. If EDTA is in the solution, it will chelate the nickel from the beads and prevent detection of the protein.

When tagging the N-terminus of a protein, the main purpose is to detect this protein by a widely commercial antibody to the tag. Many proteins that researchers intend to study do not have a specific antibody, and therefore, the utilization of a HA or FLAG tag allows easy detection. The tag is not intended to alter the native protein's function nor change its expression level in cells.

One of the purposes of biochemically tagging a protein is to facilitate its detection. As such, it is important that the tag is expressed only when the protein is expressed. Additionally, the in-frame tag is typically fused to the or -terminus of the protein to ensure minimal disruption in native protein folding and structure.

Researchers commonly use FLAG, His, HA and GST tags when biochemically tagging proteins for experiments. Often, these tags are fused in-frame to the -terminus of the protein to minimize disruption of native protein folding. Sodium dodecyl sulfate (SDS) is an anionic detergent, that when applied to proteins, destroys their folded secondary and tertiary structure. SDS is most commonly used in Western blots.

All of the answer choices are true. Creating fluorescently tagged proteins is a common laboratory practice to visualize subcellular localization of proteins. Fusion proteins are subcloned into expression vectors, therefore, it is important that the expression vector is suitable for expression in the species one wishes to express the protein. Furthermore, to minimize disruption of native protein fold, fluorescent tags are usually added to either the N or C-termini. Finally, to ensure that translation is not disrupted, the fusion must be in-frame with the native protein.

The correct answer is Northern blot. A Northern blot is used to detect or quantify RNA in a sample to measure gene expression. A biochemically tagged protein can be detected by Western blot using an antibody specific to the tag. Cell culture cells expressing a tagged protein can also be detected using an antibody specific to the tag by indirect immunofluorescence. Tagging a protein with a fluorescent protein allows researchers to track a protein's localization and movement in cells with live cell imaging. Finally, an electrophoretic mobility shift assay (EMSA) detects a protein's ability to bind DNA, but a super shift incubates the protein of interest with an antibody to determine which protein is binding DNA. In this case, a tagged protein can be probed on an EMSA.

These are all possible methods for generating a labeled DNA probe that is easier to visualize in a heterogeneous sample. In nick translation, DNase is used to nick the DNA of interests, then DNA Polymerase I replaces some nucleotides with their labeled analogues. End labeling involves the use of labeled nucleotides that prevent further elongation at the ends of the DNA molecule. PCR with a radioactive label is another method of cloning labeled DNA.

The correct answer is introduce membrane pores. Escherichia coli grows optimally at 37 degrees Celcius. However, an increase in temperature causes the plasma membrane to become more fluid, introducing small pores through which plasmid DNA can enter into the cell.

The correct answer is the ampicillin the nutrient agar plates expired. The plasmid expresses beta-lactamase, an enzyme that degrades ampicillin. Successfully transformed bacteria will be able to grow in the presence of ampicillin, but untransformed bacteria should not grow. Since the bacteria in our control transformation do not have ampicillin resistance conferred by beta-lactamase, we would expect that no growth would be observed on any nutrient agar plates with ampicillin. However, we observe uncontrolled growth on these plates, indicating that the ampicillin has expired and is no longer a viable selectable marker for transformed bacteria.

The annealing temperature is the optimal temperature at which the primers bind the template DNA sequence. This temperature takes into account several factors: the number of basepairs in the primers, the relative guanine-cytosine content, and their melting point.

None of the answer choices are correct. In order to successfully clone a bacterial gene into a mammalian expression vector, the following must be completed: First, the bacterial gene should be PCR amplified from bacterial cDNA, which contains only exons. Second, the vector backbone should be digested with restriction enzymes to linearize the vector so that the bacterial gene can be inserted at a specifc locus (multiple cloning site). Third, the amplified bacterial gene and the digested vector backbone are ligated together to create the desired expression construct.

The correct answer is cDNA. Given that this gene most likely has several introns, complementary DNA (cDNA) should be used as a template since it only contains exons. When subcloning a gene into an expression vector, only coding sequence should be present because the transcribed RNA will not be processed like it is in host cells.

Plasmids are circular pieces of DNA that are resistant to exonucleic degradation (exonucleases cut from DNA ends). Bacteria harbor native plasmids that replicate independently from their genome and express genes that often confer a survival advantage. Scientists often clone genes into plasmids and express them in a variety of host cells.

Frederick Sanger invented the chain termination method of DNA sequencing in 1977, which has been widely used since its invention for short DNA sequences. The method is know as Sanger Sequencing. Leroy Hood was responsible for automating this method, Kary Mullis was the inventor of the polymerase chain reaction to create copies of DNA, E. M. Southern lent his name to another DNA study method called Southerns, and Craig Venter was responsible for other later DNA sequencing methods.

Polymerase chain reaction requires two, not one, synthesized primer molecules to correctly amplify a DNA molecule into new copies. One primer attaches to each end of your desired DNA segment, and amplification occurs from these sites by polymerase molecules. The other options are all correct facts about PCR.

Cosmids could handle DNA insert sizes of 25-50kb. BAC's could handle insert sizes of 100-300kb, and YAC's even more. These are still used for tricky genome projects due to this large insert size, like antifreeze protein genes in Arctic fish. Plasmids can generally only handle insert sizes of 5-10kb.