Appendix 3: Additional laboratory details
Lab 3: Agarose Gel Electrophoresis
Agarose is a highly purified form of agar that is extracted from seaweed. The density of the gel is determined by the weight percentage concentration of agarose in the solution. Different concentrations of agarose are used in gels depending on the relative sizes of the fragments expected or the separation desired. Acrylamide gels can be used when small pieces of DNA are to be resolved.
The molecular weight of DNA determines its rate of migration. Typically, the smaller the molecule, the faster it will migrate through the gel. However, DNA of the same size can have differing mobilities depending upon the shape: linear, circular relaxed, or circular supercoiled. Circular supercoiled DNA migrates faster than either linear or circular relaxed DNA, while linear DNA typically migrates faster than circular relaxed DNA.
When the voltage is low, migration of DNA is proportional to the strength of the electric field, regardless of the DNA size. At higher field strengths, larger molecules move disproportionately faster so that the resolution is decreased.
The buffer system can also affect mobility. If you forget to add the running buffer to the agarose gel, electrical conductance will be decreased and the DNA will migrate very slowly. Alternatively, if you use 10x or 50x buffer to make the gel, the ionic strength will be too high, and consequently the amount of heat produced will be excessive.
The sample is loaded after the gel is just covered with buffer. It is important to add just enough buffer to cover the gel, but not a lot more. Excess buffer allows a substantial conduction pathway on top of the gel instead of the desired pathway through the gel, so migration of DNA through the gel will be slowed.
Samples are mixed with a solution containing bromphenol blue and xylene cyanol as tracking dyes; the bromphenol blue migrates about the same rate as a 0.3-0.5 kb fragment and the xylene cyanol migrates similarly to a 4-5 kb fragment. This loading dye can also contain SDS, EDTA, and sucrose. The sucrose makes the sample denser than the buffer so the sample will sink to the bottom of the well when it is loaded. Generally 1-2 μL of loading dye is added per 10 μL of sample.
Ethidium Bromide Staining
Since DNA cannot be detected in a gel, ethidium bromide, a fluorescent dye, is added to the DNA. The planar, polycyclic ethidium bromide intercalates between the bases of double-stranded DNA and RNA. Ethidium absorbs UV light at 302 nm and produces a reddish-orange fluorescence at 590 nm; the amount of this fluorescence is proportional to the mass of the DNA, assuming there is enough ethidium present to saturate all binding sites on the double stranded nucleic acid. The fluorescence of RNA is less than that of an equal mass of DNA, because most of the RNA is single-stranded and less susceptible to ethidium intercalation.
Ethidium bromide can be added to the DNA in the gel in any of three ways:
1) Ethidium bromide may be added to the gel when the hot agarose is poured onto the slide.
2) The ethidium may be added to the running buffer.
3) Following electrophoresis, the gel can be placed in a staining tray containing the dye, incubated for a short time, and then rinsed to remove any unbound ethidium bromide.
Lab 4: Transformation
When bacterial cells take up DNA they are said to undergo “transformation.” The exact mechanism of this process is unknown, but host bacterial cells must be in a state called “competence.” Healthy, mid-log phase cells are treated with a solution of ice-cold calcium chloride and then, in the presence of the recombinant DNA to be transferred, subjected to a brief heat shock. One hypothesis is that treatment of the cells at 0 ˚C crystallizes the cell membrane, stabilizing charged phosphate distribution. Then the cations in the treatment solution could complex with exposed phosphate groups to shield negative charges and a plasmid molecule might pass through “pores” in the cell’s wall/ membrane called “adhesion zones.” It is hypothesized that heat shock complements this process, possibly by creating a thermal imbalance on either side of the membrane that will move the transforming DNA through the adhesion zone into the competent host cell.
Competent cells are very fragile and must be treated with care – kept on ice, mixed very gently. The timing of the heat shock step should also be precise – if kept too long at the high temperature, the cells will die. Furthermore, careful, aseptic technique must be used throughout, because any contaminating bacterial cell will have the opportunity to grow and compete with desired cells.
After the recombinant DNA is transferred to the competent host cells, these cells are incubated in rich medium to allow them to grow and replicate the newly transferred DNA. Then aliquots of the culture are plated on a selective medium, so that only clones that received the plasmid will grow. The colonies are then screened to locate those that have the desired characteristics. Please note the difference between “selection” and “screening.” The former allows only the clones with a desired trait (antibiotic resistance in this case) to grow. The latter allows both desired and undesired to grow, but provides a way for us to differentiate among colonies with different traits.
Lab 5: Mini-Preps of Plasmids
Although all miniprep kits have slightly different procedures, the general steps are the same. SDS and NaOH are used to lyse the cells, solubilize and denature cellular constituents, begin to degrade RNA, and denature (separate the strands of) DNA. Then, in alkaline conditions, the small plasmid strands remain linked to their complementary strands. Hydrogen bonds can reform between these strands, which are close to each other when the lysate is neutralized. Proper base pairing can then occur and an intact double-stranded plasmid is reformed. Since genomic DNA will not be linked to a complementary strand and thus, not necessarily remain nearby, a rather large interlinked mass of DNA strands will result after neutralization. The potassium acetate added to neutralize the alkaline solution also elevates the salt concentration, so that the SDS and protein form a flocculent precipitate to trap the genomic DNA mass for removal by centrifugation.
Each commercial kit generally includes a proprietary resin (specially-treated glass) that is mixed with the plasmid DNA/RNA solution. The plasmid DNA sticks to the resin in a small column. The column is washed with a solution that contains ethanol, so the plasmid remains on the resin. The plasmid DNA is then eluted from the glass resin with water or TE buffer. Sometimes the DNA is stuck to the resin with high salt and eluted with low salt (water or TE). This resin replaces the alcohol precipitation steps done in other procedures and in the purification of genomic DNA.
The miniprep kit we will be using is made by Qiagen. All of the buffer formulations are proprietary, so Qiagen refers them to as P1, P2, N3 and PE.
In general the buffers do the following:
P1 – bacterial re-suspension buffer, contains RNase A to remove RNA.
P2 – alkaline lysis buffer containing NaOH-SDS. SDS causes lysis of the cell membrane. NaOH denatures chromosomal DNA, plasmid DNA, and proteins. The lysis time is optimized to release the plasmid DNA from the cell and minimize its exposure to denaturing conditions.
N3 – neutralization buffer containing acidic potassium acetate. High salt in the buffer precipitates the detergent and traps proteins and chromosomal DNA in salt-detergent complexes. The small, covalently closed plasmid DNA renatures and remains in solution. This high salt buffer also allows the plasmid DNA to bind to the resin in the spin columns.
PE – wash buffer that removes salts from the column-bound DNA.
Lab 7: Gel Electrophoresis
Gel electrophoresis is a powerful separation technique used for analyzing many biologically important molecules, including both proteins and nucleic acids. Since most biological polymers carry a net charge in solution, they will migrate in an electric field. The molecular migration is stabilized by polyacrylamide, agarose, or some other inert, solid support.
Electrophoretic mobility can then be used to distinguish molecules that differ only slightly in net charge or shape, to detect amino acid residue changes in proteins, to determine protein molecular masses, or the size of polynucleotide fragments. This laboratory exercise focuses on protein electrophoresis.
The function of sodium dodecyl sulfate (SDS) and β-mercaptoethanol (BME) in denaturing gel electrophoresis
An important application of protein electrophoresis requires the use of the denaturing detergent sodium dodecyl sulfate (SDS) and a reducing agent. At neutral pH in 1% SDS and 0.1 M β-mercaptoethanol (BME), most protein molecules unfold, and inter- and intra-chain disulfide linkages are reduced. All higher order protein structure is therefore lost, and multi-subunit proteins separate into their constituent subunits. Each protein chain is coated with a layer of SDS molecules such that negatively charged sulfate groups of the detergent, which usually far outnumber the charged groups belonging to the protein side chains, are exposed to the aqueous medium.
Hence, the presence of SDS confers a large negative charge to the protein-detergent complex. Moreover, SDS is bound to most proteins in a nearly constant ratio (about 1.4 g SDS per g protein), which provides a nearly constant charge to mass ratio. Consequently, proteins treated with SDS and β-mercaptoethanol and then subjected to electrophoresis migrate toward the anode at a rate that is determined by the mass of the polypeptide, the porosity of the gel, and the electric field potential. In general, a small molecular mass polypeptide will have a faster rate of migration than a larger one. This observation forms the basis for using SDS polyacrylamide gels to characterize mixtures of proteins based on molecular mass.
Gels of a given porosity (achieved by adjusting the acrylamide and crosslinker concentrations) are calibrated with standard proteins of known molecular masses. The molecular mass of an unknown polypeptide can then be estimated from a standard curve for the given gel system.
Discontinuous gel electrophoresis
An important refinement of electrophoresis is the method of discontinuous gel electrophoresis. A discontinuous gel system consists of two acrylamide gels; one gel is cast directly on top of the other. The lower gel (which is poured first and allowed to polymerize) is called the resolving gel. The resolving gel is so called because resolution takes place in this gel on the basis of size (due to sieving action of the polyacrylamide). The resolving gel contains an acrylamide concentration ranging from 5-30%, depending on the range of protein molecular masses to be separated. The higher the percentage of acrylamide used, the lower the molecular weight range that will be resolved.
The upper gel, called a stacking gel, contains a much lower percentage of acrylamide than the resolving gel (usually about 2-3%), and offers little separation based on molecular mass. Instead, the stacking gel compresses the protein sample into a narrow band before it enters the resolving gel and thereby improves the resolution of the resolving gel. The stacking phenomenon is provided by differences in the pH and ionic strength of the buffers in the stacking gel, the resolving gel, and the buffer reservoir. The larger the volume of the sample to be loaded, the more useful a stacking gel becomes.
The stacking phenomenon
The protein bands are stacked as follows: At the pH of the stacking gel (6.8), the glycine amino group is predominantly in the protonated form, so that the amino acid is an electronically neutral zwitterion. In contrast, the protein coated with SDS has a net negative charge. In this state, protein has an electrophoretic mobility between that of the glycine zwitterion (not migrating) and Cl-(the fast-moving counter-ion in the discontinuous gel buffer). The Cl-ion migrates rapidly primarily because it is small and is followed by SDS-coated protein (having a large size, but also a large net charge), followed by glycine (which has low net charge due to the pH equilibrium). As the Cl-ion moves ahead of the glycine, a drop in conductivity occurs at the interface between the Cl-ions and the glycine ions. This decrease in conductivity gives rise to a voltage gradient that accelerates the slower moving protein and glycine. Ions ahead of the glycine-chloride interface experience are not exposed to a voltage gradient because the conductivity is not affected in this region: therefore, they are not accelerated. By continued acceleration into a zone of high conductivity, the protein band, which sits between the chloride-glycine interface, is stacked into a sharp band. The formation of a sharp band greatly improves the resolution that can be attained in the resolving gel.