It is common nowadays to go to the doctor’s office for a sore throat or a host of other ailments and walk out with a prescription for antibiotics. We hear time and again that this over use of antibiotics in western society must have negative consequences. Indeed, the overuse and misuse of antibiotics have led to many cases in which microorganisms survive an encounter with a drug. This can lead to genetic mutations in the bacteria that result in resistance to that drug. Furthermore, this problem is amplified because these resistant bacterial strains are known to be able to transfer genes that result in resistance to other bacteria as well. This can even give rise to some bacteria developing resistance to multiple antibacterial drugs; such bacteria are known as multidrug resistant (MDR) bacteria or colloquially, super bacteria or super bugs. This leads to the need for the development of new, effective antibacterial drugs. Thus we have developed a bioassay to find chemical compounds that specifically target the FabG protein involved in the fatty acid synthase (FAS) pathway in Escherichia coli.
We have chosen to look for antibiotics that target the fatty acid biosynthesis pathway for several reasons: first, because it is an appealing but still largely unutilized target for the development of new antibacterial compounds; secondly, because bacteria use a different type of enzyme (type II FAS) than mammals (type I FAS) to synthesize fatty acids. These two elements mean that there is room for discovery of new compounds that will inhibit this essential pathway but at the same time not adversely affect the fatty acid biosynthesis pathway in humans. Many bacteria use a series of type II FAS enzymes named FabX proteins (where X = A, B, C, D, etc.). FabF, FabH, and FabI have all been put forward as antibacterial drug targets. The related enzyme FabG is also implicated in this series as a good drug target but has not yet been widely explored; thus we have set out to explore FabG as the target of this drug discovery bioassay.1)
The FabG enzyme is involved in the elongation cycle of the fatty acid biosynthesis pathway in E. coli. It catalyzes the reduction with the help of the coenzyme NADH of a carbonyl group to an alcohol group from one intermediate to the next. This bioassay will hopefully result in the discovery of a compound that inhibits the FabG enzyme in this reaction step of the fatty acid biosynthesis pathway.
by Jess Coulter
A primer is a short synthetic oligonucleotide that is used in molecular techniques such as PCR and DNA sequencing. The primers are designed to have a sequence which is the reverse complement of a region of template or target DNA to which we wish the primer to stick to. The primer is used to amplify the fabG gene from E. coli DNA.
Length: 18-23 bases
GC%: 50-60% so it is able to stick to the DNA
TM: 55°-65° C (TM of Forward and Reverse primers should be within 3° C of each other)
The primers should not have self-complimentary sequences or mismatches with genomic sequences that are not of interest
To allow directional cloning, the forward PCR primer must contain the sequence “CACC” at the 5’ end of the primer. These four nucleotides base pair with the overhang sequence, “GTGG,” in each pET TOPO vector.
In order for the PCR product to clone directionally, the reverse PCR primer must not be complementary to the overhang sequence GTGG at the 5’ end.
The N-terminus of the protein is encoded by:
1. The forward PCR primer should be:
5’ – CACCATGGCCCCCCCGACCGAT – 3’
2. For the reverse primer, analyze the C-terminus of the protein:
…GCG GTT AAG TCG GAG CAC TCG ACG ACT GCA TAG – 3’
3. The reverse PCR primer should be:
5’ – CTA TGC AGT CGT CGA GTG CTC CGA CTT – 3’
In our experiment, we used the NCBI GenBank to find the sequence of the fabG gene in E. coli. From this gene, we created forward and reverse primer sequences based on the requirements above, but all of our sequences turned out to be reversed from what they should have been. Dr. Joyner went through each of our sequences and fixed them for us. These were the final primer sequences:
Polymerase Chain Reaction (PCR) is a versatile lab technique used to amplify a specific portion of a DNA sequence. The typical PCR system contains the following components:
• Template DNA: Collection of DNA that contains the desired sequence for amplification
• DNA polymerase: This must be a polymerase that operates optimally at a temperature around 70 °C due to the high temperatures used in the PCR protocol. Usually, Taq (Thermus aquaticus) polymerase is chosen.
• Nucleotides: These are the building-blocks that the DNA polymerase uses to synthesize a new DNA strand.
• Salts: This includes monovalent and divalent cations. These are used to mimic cell conditions.
• Buffer Solution: This provides an adequate chemical environment for maximal activity and stability of the DNA polymerase used.
• Primers: Primers are used that are complementary to the sense and anti-sense strand of the DNA target to be amplified. Thus, both a forward and a reverse primer are utilized.
All of the above-mentioned ingredients are added to a reaction vessel and loaded into a thermal cycler.
A general PCR protocol consists of three standard phases.
Melt Phase: In this phase, the thermal cycler heats the reaction mixture to a temperature of around 95 °C for 30 seconds. This causes the DNA template to melt, yielding single-stranded DNA molecules.
Annealing Phase: In this phase, the temperature of the system is lowered to 55 °C and held for 30 seconds. During this step, the forward and reverse primers anneal to the corresponding sequences of the single-stranded DNA template. An optimal annealing temperature is generally about 3-5 °C less than the melting temperature (Tm) of the primers used. Hydrogen bonds form between the primer sequence and DNA template, and the chosen polymerase binds to the template-primer hybrid and initiates DNA synthesis.
Extension Phase: In this phase, the temperature is raised to 72 °C and held for 1 minute. In this step, the DNA polymerase used synthesizes a new DNA strand complementary to the DNA template by using nucleotides present in solution, extending the strand in the 5’ to 3’ direction.
If n represents the number of cycles performed in a given PCR system, then PCR will optimally produce 2n copies of the desired DNA sequence.
PCR was utilized in order to target and amplify the sequence of DNA coding for the fabG protein. The principles of primer design were used to select successful primers for hybridization to the fabG gene sequence.
Phase 1 (Melt): 30 seconds at 95 °C
Phase 2 (Anneal): 30 seconds at 54 °C
Phase 3 (Extend): 1 minute at 72 °C
Amount of Cycles: 30
The reaction tubes and plates were put on ice prior to the PCR being run. The following components were added to the PCR reaction vessel:
The contents of the reaction vessel were mixed and placed on ice. Then, the reaction vessel was capped and loaded into the thermal cycler. The program was then initiated and run according to the parameters outlined above. Purity of the PCR product was assessed using agarose gel electrophoresis. A bright band near 730 base pairs (length of the fabG gene) was visualized and thus confirmed successful amplification of the gene sequence. The purity of the PCR product was further assessed using a NanoDrop UV-Vis Spectrophotometer. The ratio of absorbances at 260 and 280 nm was calculated; PCR products were considered “pure” if this A260 / A280 ratio was greater than or equal to 1.80.
DNA extraction is a procedure used to collect DNA for further analysis. In this case, E. coli DNA was extracted for further analysis of the fabG gene present in the cells’ DNA. In this extraction, there were six basic steps.
The purpose of the DNA extraction step was to extract DNA from E. coli using a DNA spin column kit (Geneaid DNA Extraction Kit). This step was necessary for the bioassay for the extracted DNA was used for amplification of the fabG gene and the integrity. To verify that the DNA extraction was successful, the purity of the extracted DNA was determined through agarose gel electrophoreseis and UV-Vis spectoscopy.
Cultured bacterial cells of a concentration up to 1×10<sup>9<sup> were transferred to a 1.5mL centrifuge tube. Samples were centrifuged for 1 minutes at 14,000-16,000g, and the supernatant was discarded. The cell pellet was re-suspended with 200uL of GT buffer. Samples were incubated at room temperature for 5 minutes.
To the sample mix, 200 uL of GB buffer was added and mixed through vigorous shaking for 5 seconds. The sample was incubated at 60 degrees C for 10 minutes to ensure that the sample lysate was clear.
To the clear lysate, 200 uL of absolute ethanol was added. A GD column was placed in a 2 mL collection tube, and the entire sample mixture was added to the column. The sample was centrifuged at 14,000-16,000xg for 2 minutes, and the flow through was discarded.
To the GD column containing the extracted DNA, 400 uL of WI buffer was added, and the column was centrifuged at 14,000-16,00xg for 30 seconds. The flow through was once again discarded. To the column, 600uL of wash buffer with ethanol was added, and the sample was centrifuged at 14,000-16,000xg for 30 seconds. The flow through was discarded once more. The column was centrifuged again for 3 minutes at 14,000-16,000xg to dry the column matrix.
Standard elution volume is 100uL. If the sample to be used is less, reduce the elution volume to 30-50uL to increase the DNA concentration. If higher DNA yield is required, repeat the DNA elution step to increase DNA recovery until the total elution volume is 200uL. The dried GD column was transferred to a clean 1.5mL microcentrifuge tube, and 100uL of pre-heated elution buffer or TE buffer was added to the center of the column matrix. The tube was left to stand for at least 3 minutes to ensure that the elution buffer was absorbed by the matrix. The column and microcentrifuge tube was centrifuged at 14,000-16,000xg for 30 seconds to elute the purified DNA.
The purified DNA was collected and stored for further analysis. More specifically, the extracted DNA was amplified using PCR. As previously described, PCR is a method used to amplify specific sequences of DNA. In this case, PCR was used to amplify the fabG gene sequence in the purified E. coli DNA. Once PCR was performed, the purity of the DNA was assessed through UV-Vis spectroscopy. An agarose gel was also run in order to verify the purity as well as the presence of amplified concentrations of the fabG gene.
by Cameron Kubota and Daniel Rossie
After obtaining the fabG gene from the PCR reaction, the gene had to be cloned and overexpressed in a new E. Coli DNA. In order to successfully implant the gene into another E. Coli cell, we followed a cloning and transformation protocol as laid out in the Invitrogen Champion™ pET Directional TOPO® Expression Kits Manual. Circular DNA molecules called plasmids facilitate this introduction of the gene into a new cell.
Plasmids are used as a vehicle to transfer replicated genes in order to overexpress the gene product in a new host cell. Plasmid DNA is transformed into new cells in order to overexpress a target gene, which in our study was the fabG gene isolated from E. Coli DNA. DNA extracted from E. Coli and replicated through PCR was inserted directly into our pET100/D-TOPO vector. According to the suggestions of Invitrogen, a .5:1 to 2:1 molar ratio of PCR product to vector should be used in order to obtain the highest cloning efficiency possible.
The general method for inserting the targeted gene into a plasmid is by introducing an overhang on both ends of the gene that are complementary to the overhangs on the vector. Restriction enzymes are used to cut a gap in the plasmid, leaving behind a series of about four unpaired bases on both sides of the gap. The four complementary bases are introduced onto both ends of the targeted gene creating “sticky” ends, or ends that will readily form hydrogen bonds and complete the plasmid sequence. However, the pET Topo kit utilizes an overhang on one end, but a blunt-end method on the other. The DNA Ligase then zips up the plasmid to form a complete circular vector.
We chose the three PCR product solutions that contained the highest DNA concentrations. Two replicates of each PCR product were carried out by adding 1.1 μL of the PCR product, 1 μL of the pET Topo kit salt solution, and 1 μL of the vector to a clean vial. To bring the total mixture volume to 6 μL, 2.9 μL of sterile water was added. Three more samples were made by adding the same volumes of each component but replacing the 1.1 μL of the PCR product with 1.1 μL of the positive control sample. The samples were gently mixed and incubated for 5 minutes at room temperature and then transferred to an ice bath.
The goal of bacterial transformation is to implant another set of DNA to be replicated within the new cell. For our study, we cloned the fabG gene into plasmid DNA and then transformed the plasmid into E. Coli cells in order to overexpress the fabG gene product for further bioassay studies. Overexpression of the fabG gene in a new E. Coli cell allows us to target the fabG gene specifically for antibacterial bioassays.
3 μL of the cloned vector was added to the vial of OneShot TOP10 Chemically Competent E. Coli. This mixture was incubated on ice for 30 minutes and then heat-shocked for 30 seconds at 42º C without shaking. Then, the sample was transferred immediately to ice and 250 μL of room temperature S.O.C. medium was added. The vial was then placed in the shaking incubator and was shaken at 200 rpm for one hour at 37º C.
After an hour in the shaking incubator, the plasmid should be completely transformed into the new E. Coli cells. The solution containing these cells was spread on a pre-warmed selective agar plate containing Ampicillin and incubated overnight at 37º C. Because our vector has an Ampicillin resistant region, the plate used for streaking was selective for Ampicillin resistant cells, killing off any other bacteria that could have been introduced into the sample.
Following the purification and isolation of the plasmid DNA, it is necessary to verify that the correct DNA sequence was properly incorporated into the plasmid. This verification was performed using restriction enzyme digestion, a process of using restriction enzymes to cut at various sites on the plasmid. The results of this digest were analyzed using agarose gel electrophoresis.
Restriction enzymes are enzymes isolated from bacteria that are able to cut at specific sequences on DNA strands to form restriction fragments. Many restriction enzymes have been studied and are available commercially. They can therefore be used to cut at specific locations within various gene sequences, called restriction sites. For this analysis, the enzymes AluI, PvuI, and MseI were available for use. AluI and MseI both cut at 25 different sites within the plasmid while PvuI only cuts at one location within the plasmid. The entire plasmid length is 5,764 base pairs, while the fabG promoter sequence is 730-750 base pairs long.
Once the restriction enzyme had been chosen, its restriction sites were analyzed to determine where it would be likely to cut in our plasmid. From the restriction sites, the lengths of several likely restriction fragments were calculated for use in the analysis of the gel electrophoresis experiment. By predicting likely lengths of restriction fragments, it is possible to determine whether or not the fabG promoter sequence is properly incorporated and oriented in the plasmid.
Trials were performed using all three restriction enzymes, using one unit of enzyme, or the amount required to digest one μg of DNA in one hour at 37°C. 10μL of plasmid DNA were combined with 2μL of restriction enzyme, an excess amount. The final volume was then brought to 50μL with buffer and allowed to digest at 37°C for one hour. Following the digestion period, the samples were run on an agarose gel electrophoresis to determine whether DNA restriction fragments of the expected length were obtained.
After allowing the restriction enzyme solution to incubate for one hour, the concentration of plasmid DNA was assessed using a nanodrop spectrophotometer. 2 µL of each student’s digest was placed on the sample-loading detector in order to determine the sample concentrations. The following concentrations were obtained:
The concentrations of plasmid DNA were not ideal for running a successful gel electrophoresis. The low concentrations indicated that the plasmid DNA purification protocol failed at some point. Those who obtained 0% percent concentration (or less) of plasmid DNA in their final solutions investigated several of the earlier fractions saved from different points in the restriction enzyme digest/plasmid DNA purification protocol. The results from this investigation indicated that most of the plasmid DNA was lost in the third fraction.
Despite the disappointingly low concentrations of plasmid DNA, it was decided to go forward with the gel electrophoresis in the hopes that meaningful results could still be obtained in determining whether or not the fabG gene was cloned correctly into the plasmid DNA.
It was decided to use a 1.5% concentration of agarose gel to view the separation of DNA fragments. The agarose gel was made using the following procedure:
Once the agarose gels were made, 10µL of each student’s plasmid DNA samples were put into solution with 20µL of loading buffer. Each student then used a micropipette to load 25µL of their solution into one of the agarose wells. A DNA ladder was loaded into one of the wells as a reference. The gel was run at 100V and 80mA .
The results failed to verify whether the fabG gene had been cloned into the plasmid correctly. Although this could not be verified, it was decided to continue with the bioassay under the assumption that the gene had been cloned into the E. Coli correctly because of time constraints that would not allow us to completely restart the experiment and continue on to the drug discovery portion of the semester.