Abstract:
An unknown segment of DNA was recovered from Bactrocera dorsalis and subjected to characterization using insertion into vectors (EMBL3 and pUC19), restriction enzyme mapping, Southern blot sequence verification, and PCR amplification. The genomic library was created successfully with 45 plaques available for analysis. The DNA was positively inserted into the pUC19 vector, as verified by the presence of white colonies on the selective media (LB-amp-Xgal). A restriction enzyme map illustrated that the insert contained recognition sites for HindIII 500 bp, Xho1 1000 bp, and Pst1 3500 bp from the beginning of the insert. The Southern blot verified the presence of the target sequence in the linearized recombinant plasmid. The 720-bp sequence was also analyzed using database analysis, and primers were designed for the amplification reaction. PCR revealed a fragment size of between 100 and 200 base pairs, indicating that the primers successfully isolated the 161-bp target sequence. Following these tests, the unknown segment was found to contain an actin-coding gene. Actin is highly conserved among species. It functions in the eukaryotic cytoskeleton and in muscular contraction.
Wednesday, December 13, 2006
Report Introduction
Introduction:
The analysis of DNA samples taken from organism allows a better understanding of the function and structure of that organism. DNA fragments can be sequenced and the results recorded in computer databases. Bioinformatic analysis of future experimentally obtained DNA sequences can be compared to known sequences. This aids in taxonomic organization, when comparing related species and establishing evolutionary patterns based on genomic analysis.
DNA analysis functions in this way due to the highly conserved nature of DNA and DNA’s role as the genetic origin of all cellular form and function. The sequence of nucleotide base pairs of DNA, which can be 109 bp or larger, changes over time and generations, but only very slowly. Mutations occur, but the rate is very slow, and often mutations do not provide a survival advantage to the individual, and are not conserved.
Many techniques have been developed that use molecular biological and genetic knowledge. Amplification of a DNA is often necessary for further analysis. This can be accomplished in vivo or in vitro. In vivo amplification requires the use of a vector, such as a plasmid or a virus, and host cells. The DNA fragment is ligated into the vector, which makes its way into the host cells. Bacteria are used as host cells due to their high rates of population growth. Once the fragment and vector are amplified using the replication machinery of the host cell, the DNA can be recovered. In vitro amplification (polymerase chain reaction, PCR) utilizes a mixture of the raw ingredients of DNA replication in optimal pH conditions. Temperature changes are used to denature, anneal, and extend the DNA. The key to PCR is the use of Taq (from Thermus aquaticus) polymerase, which is not degraded by high temperatures.
Blots and probes can verify the presence of specific DNA sequences. A probe is a marker-containing sequence complimentary to the target sequence. The probe will hybridize to the target sequence if the target is present. Hybridization can be verified by the presence of the marker. If hybridization does not occur, the probe will be washed off, and the marker will not be observed.
Studies using fruit flies have given significant credibility to theories involving allele distribution and genetic coding of traits. Fruit flies are useful organisms for the study of genetics for several reasons. They are small and easy to grow in laboratory settings. They have a short generation time (around 2 weeks), and females often produce many offspring, allowing greater statistical significance in calculated ratios used for allele distribution. Males and females are easily discernable, facilitating the mating process. Also, fruit flies have only four pairs of chromosomes (3 autosomal, 1 sex), and the genomes of some species, such as Drosophila melanogaster, have been entirely sequenced.
Actin is a globular structural protein that forms in an actin filament, which form the cytoskeleton (structural support) of eukaryotic cells. Actin also combines with myosin form actomyosin in muscle cells, which uses ATP as an energy source for muscle contraction. Actin-coding genes are some of the most highly conserved genes among different species. For example, the actin gene of Homo sapiens is over 80% similar to the actin gene of Saccharomyces cerevisiae, a budding yeast.
In this experiment, our objective was to isolate a fragment of genomic DNA from Bactrocera dorsalis and characterize and sequence this segment, noting the function of genes present in the region.
The analysis of DNA samples taken from organism allows a better understanding of the function and structure of that organism. DNA fragments can be sequenced and the results recorded in computer databases. Bioinformatic analysis of future experimentally obtained DNA sequences can be compared to known sequences. This aids in taxonomic organization, when comparing related species and establishing evolutionary patterns based on genomic analysis.
DNA analysis functions in this way due to the highly conserved nature of DNA and DNA’s role as the genetic origin of all cellular form and function. The sequence of nucleotide base pairs of DNA, which can be 109 bp or larger, changes over time and generations, but only very slowly. Mutations occur, but the rate is very slow, and often mutations do not provide a survival advantage to the individual, and are not conserved.
Many techniques have been developed that use molecular biological and genetic knowledge. Amplification of a DNA is often necessary for further analysis. This can be accomplished in vivo or in vitro. In vivo amplification requires the use of a vector, such as a plasmid or a virus, and host cells. The DNA fragment is ligated into the vector, which makes its way into the host cells. Bacteria are used as host cells due to their high rates of population growth. Once the fragment and vector are amplified using the replication machinery of the host cell, the DNA can be recovered. In vitro amplification (polymerase chain reaction, PCR) utilizes a mixture of the raw ingredients of DNA replication in optimal pH conditions. Temperature changes are used to denature, anneal, and extend the DNA. The key to PCR is the use of Taq (from Thermus aquaticus) polymerase, which is not degraded by high temperatures.
Blots and probes can verify the presence of specific DNA sequences. A probe is a marker-containing sequence complimentary to the target sequence. The probe will hybridize to the target sequence if the target is present. Hybridization can be verified by the presence of the marker. If hybridization does not occur, the probe will be washed off, and the marker will not be observed.
Studies using fruit flies have given significant credibility to theories involving allele distribution and genetic coding of traits. Fruit flies are useful organisms for the study of genetics for several reasons. They are small and easy to grow in laboratory settings. They have a short generation time (around 2 weeks), and females often produce many offspring, allowing greater statistical significance in calculated ratios used for allele distribution. Males and females are easily discernable, facilitating the mating process. Also, fruit flies have only four pairs of chromosomes (3 autosomal, 1 sex), and the genomes of some species, such as Drosophila melanogaster, have been entirely sequenced.
Actin is a globular structural protein that forms in an actin filament, which form the cytoskeleton (structural support) of eukaryotic cells. Actin also combines with myosin form actomyosin in muscle cells, which uses ATP as an energy source for muscle contraction. Actin-coding genes are some of the most highly conserved genes among different species. For example, the actin gene of Homo sapiens is over 80% similar to the actin gene of Saccharomyces cerevisiae, a budding yeast.
In this experiment, our objective was to isolate a fragment of genomic DNA from Bactrocera dorsalis and characterize and sequence this segment, noting the function of genes present in the region.
Report Procedure
Materials and Methods:
Previously, isolated genomic DNA from Bactrocera dorsalis was treated with the restriction enzyme EcoRI, and the resultant fragments were ligated into the EMBL3 enterobacteria phage λ vector.
5 μl of this λ-phage solution and 95 μl buffer (NaCl and Mg2+) were added to 200 μl of Escherichia coli host culture suspended in MgSO4. The sample was incubated at 37ºC for 20 minutes. Following incubation, the sample was mixed with 50ºC warm top agar and plated onto fresh nutrient agar. After hardening, the plate culture was incubated at 37ºC for 72 hours.
One plaque from the resultant genome library was selected, and the recombinant DNA (EMBL3 backbone with B. dorsalis genomic DNA insert) was recovered, amplified, and treated with EcoRI to isolate the genomic DNA insert segment. Additionally, 2 μl of pUC19 plasmid vector was treated with EcoRI. 2 μl (100 ng) of isolated genomic DNA insert was ligated into 2 μl (100 ng) pUC19 plasmid vector using 1 μl of DNA ligase enzymes, in 2 μl buffer and 14 μl distilled water. The mixture was incubated at 20ºC for 5 minutes, and then chilled on ice. 10 μl of the reaction mixture was added to 40 μl of competent E. coli cells. The cells were incubated on ice for 30 minutes, heat shocked for 40 seconds in a 37ºC water bath, and returned to ice incubation for 2 minutes. Following incubation, 950 μl of LB broth was added, and the culture was incubated for 35 minutes at 37ºC. Following incubation, 75 μl of the cell culture was plated on selective solid LB agar containing Xgal and ampicillin and incubated for 24 hours at 37ºC.
Following incubation, one colony was selected to create a restriction enzyme map of the recombinant DNA plasmid. Five single enzyme digests and five double enzyme digests were performed on the recombinant DNA plasmid (pUC19 plasmid vector and the insert DNA from Bactrocera dorsalis). For the single enzyme digests, 1 μl of the restriction enzymes BamHI, HindIII, Pst1, EcoRI, and Xho1 each were added to separate preparations containing 5 μl recombinant plasmid DNA, 2 μl buffer solution, and 12 μl distilled H2O. For the double enzyme digests, five enzyme combinations were selected: EcoRI with Xho1, HindIII with Pst1, Xho1 with Pst1, Eco RI with HindIII, and EcoRI with Pst1. 1 μl of each enzyme was added to their respective preparations containing 5 μl of recombinant plasmid DNA, 2 μl of buffer solution, and 11 μl distilled H2O. A control containing only EcoRI and the pUC19 plasmid vector was also prepared. The 11 samples were incubated and electophoresed for 30 minutes at 90 volts in 1.0% agarose gel containing ethidium bromide.
A Southern blot of the recombinant plasmid DNA was prepared. Two samples of DNA (5 µl), the recombinant plasmid DNA (pUC19 with Bactrocera dorsalis genomic DNA insert) and a negative control (pUC19 without insert), were linearized by treatment with the restriction enzyme BamHI. The samples were denatured with 10 µl of denaturing solution (0.3 M NaOH) and incubated at 65º C for 30 minutes. Following incubation, each sample (15 µl) was dotted onto a nylon membrane and cross-linked with UV radiation for 2 minutes. After irradiation, the membrane was placed into a sealable plastic bag. 5 ml of prehybridization solution and the denatured DIG-UTP probe were added, and the sample was incubated for 24 hours. After incubation, the membrane was washed for two 5-minute cycles with 2xSSC, 0.1% SDS at 20ºC, and two 10-minute cycles with 0.5xSSC, 0.1% SDS at 55ºC. The membrane was rinsed in buffer 1 for one minute, and then incubated in a blocking solution, for 20 minutes. The membrane was placed in a bag containing enzyme-conjugated (alkaline phosphatase) anti-DIG antibody solution, and incubated at 20ºC for 25 minutes with agitation. The membrane was washed for two 10-minute cycles with buffer 1. Buffer 3 was added for two minutes and then discarded. The color detection (NBT) solution was added, and the membrane was incubated in darkness at 20ºC for 15 minutes.
A sample of the recombinant plasmid was treated with Pst1 restriction enzyme creating two linear segments. The segment containing only B. dorsalis DNA was sequenced up to 720 base pairs. Primers selected for this sequence, using Primer3 (http://frodo.wi.mit.edu/) design software, were ACTGCCAAAATGTGTGACGA (forward primer) and TTCTGACCCATACCCACCAT (reverse primer). 0.5 µl of the 720-bp segment of B. dorsalis DNA was added to 24.5 µl of PCR amplification reaction (rxn) mixture, which contained 2.5 µl PCR buffer (10X), 17.75 µl distilled H2O, 0.5 µl dNTP mix, 2.5 µl MgCl2 (1.5 mM), 0.5 µl of forward primer (20 pmol), 0.5 µl of reverse primer (20 pmol), and 0.25 µl of Taq polymerase. A negative control was also created by adding 0.5 µl H2O to another vial containing 24.5 µl of PCR rxn mix (noted above). The experimental sample and the control sample were placed into the BioRad Thermocycler, which produced 30 cycles of the following temperature sequence: 1-minute denaturation stage at 94ºC, 30-second annealing stage at 55ºC, 1-minute extension stage at 72ºC. The initial denaturation stage was 2 minutes at 94ºC, and the final extension stage was 7 minutes at 72ºC. Following amplification, both samples were electrophoresed in 2% agarose gel containing ethidium bromide for 30 minutes at 90 volts.
Previously, isolated genomic DNA from Bactrocera dorsalis was treated with the restriction enzyme EcoRI, and the resultant fragments were ligated into the EMBL3 enterobacteria phage λ vector.
5 μl of this λ-phage solution and 95 μl buffer (NaCl and Mg2+) were added to 200 μl of Escherichia coli host culture suspended in MgSO4. The sample was incubated at 37ºC for 20 minutes. Following incubation, the sample was mixed with 50ºC warm top agar and plated onto fresh nutrient agar. After hardening, the plate culture was incubated at 37ºC for 72 hours.
One plaque from the resultant genome library was selected, and the recombinant DNA (EMBL3 backbone with B. dorsalis genomic DNA insert) was recovered, amplified, and treated with EcoRI to isolate the genomic DNA insert segment. Additionally, 2 μl of pUC19 plasmid vector was treated with EcoRI. 2 μl (100 ng) of isolated genomic DNA insert was ligated into 2 μl (100 ng) pUC19 plasmid vector using 1 μl of DNA ligase enzymes, in 2 μl buffer and 14 μl distilled water. The mixture was incubated at 20ºC for 5 minutes, and then chilled on ice. 10 μl of the reaction mixture was added to 40 μl of competent E. coli cells. The cells were incubated on ice for 30 minutes, heat shocked for 40 seconds in a 37ºC water bath, and returned to ice incubation for 2 minutes. Following incubation, 950 μl of LB broth was added, and the culture was incubated for 35 minutes at 37ºC. Following incubation, 75 μl of the cell culture was plated on selective solid LB agar containing Xgal and ampicillin and incubated for 24 hours at 37ºC.
Following incubation, one colony was selected to create a restriction enzyme map of the recombinant DNA plasmid. Five single enzyme digests and five double enzyme digests were performed on the recombinant DNA plasmid (pUC19 plasmid vector and the insert DNA from Bactrocera dorsalis). For the single enzyme digests, 1 μl of the restriction enzymes BamHI, HindIII, Pst1, EcoRI, and Xho1 each were added to separate preparations containing 5 μl recombinant plasmid DNA, 2 μl buffer solution, and 12 μl distilled H2O. For the double enzyme digests, five enzyme combinations were selected: EcoRI with Xho1, HindIII with Pst1, Xho1 with Pst1, Eco RI with HindIII, and EcoRI with Pst1. 1 μl of each enzyme was added to their respective preparations containing 5 μl of recombinant plasmid DNA, 2 μl of buffer solution, and 11 μl distilled H2O. A control containing only EcoRI and the pUC19 plasmid vector was also prepared. The 11 samples were incubated and electophoresed for 30 minutes at 90 volts in 1.0% agarose gel containing ethidium bromide.
A Southern blot of the recombinant plasmid DNA was prepared. Two samples of DNA (5 µl), the recombinant plasmid DNA (pUC19 with Bactrocera dorsalis genomic DNA insert) and a negative control (pUC19 without insert), were linearized by treatment with the restriction enzyme BamHI. The samples were denatured with 10 µl of denaturing solution (0.3 M NaOH) and incubated at 65º C for 30 minutes. Following incubation, each sample (15 µl) was dotted onto a nylon membrane and cross-linked with UV radiation for 2 minutes. After irradiation, the membrane was placed into a sealable plastic bag. 5 ml of prehybridization solution and the denatured DIG-UTP probe were added, and the sample was incubated for 24 hours. After incubation, the membrane was washed for two 5-minute cycles with 2xSSC, 0.1% SDS at 20ºC, and two 10-minute cycles with 0.5xSSC, 0.1% SDS at 55ºC. The membrane was rinsed in buffer 1 for one minute, and then incubated in a blocking solution, for 20 minutes. The membrane was placed in a bag containing enzyme-conjugated (alkaline phosphatase) anti-DIG antibody solution, and incubated at 20ºC for 25 minutes with agitation. The membrane was washed for two 10-minute cycles with buffer 1. Buffer 3 was added for two minutes and then discarded. The color detection (NBT) solution was added, and the membrane was incubated in darkness at 20ºC for 15 minutes.
A sample of the recombinant plasmid was treated with Pst1 restriction enzyme creating two linear segments. The segment containing only B. dorsalis DNA was sequenced up to 720 base pairs. Primers selected for this sequence, using Primer3 (http://frodo.wi.mit.edu/) design software, were ACTGCCAAAATGTGTGACGA (forward primer) and TTCTGACCCATACCCACCAT (reverse primer). 0.5 µl of the 720-bp segment of B. dorsalis DNA was added to 24.5 µl of PCR amplification reaction (rxn) mixture, which contained 2.5 µl PCR buffer (10X), 17.75 µl distilled H2O, 0.5 µl dNTP mix, 2.5 µl MgCl2 (1.5 mM), 0.5 µl of forward primer (20 pmol), 0.5 µl of reverse primer (20 pmol), and 0.25 µl of Taq polymerase. A negative control was also created by adding 0.5 µl H2O to another vial containing 24.5 µl of PCR rxn mix (noted above). The experimental sample and the control sample were placed into the BioRad Thermocycler, which produced 30 cycles of the following temperature sequence: 1-minute denaturation stage at 94ºC, 30-second annealing stage at 55ºC, 1-minute extension stage at 72ºC. The initial denaturation stage was 2 minutes at 94ºC, and the final extension stage was 7 minutes at 72ºC. Following amplification, both samples were electrophoresed in 2% agarose gel containing ethidium bromide for 30 minutes at 90 volts.
Report Results
Results:
A genome library was created on the plate containing Escherichia coli cells infected with λ-phage containing inserts of genomic Bactrocera dorsalis DNA. A bacterial lawn covered the agar surface, and 45 plaques were observed, as well as two short fingers of no bacterial growth (see Figure 1a).
The plate of cultured E. coli cells containing recombinant DNA plasmids (pUC19 vector with ligated B. dorsalis DNA) produced 476 small circular blue colonies and 512 small circular white colonies. Often the colonies were confluent (Figure 1c).
The agarose gel electrophoresis containing restriction-enzyme-digested DNA samples contained many bands (Figure 2). Lanes 1 and 13 contained 1-kbp DNA marker. Lane 2 (control, pUC19 vector with EcoR1) showed a band at Rf 0.325, lane 3 (BamHI), lane 4 (HindIII), and lane 7 (XhoI) showed bands at Rf 0.125. Lane 5 (Pst1) showed bands at Rf 0.20 and 0.26. Lane 6 (EcoR1) showed bands at Rf 0.175 and 0.325. Lane 8 (EcoR1, XhoI) showed bands at Rf 0.20, 0.325, and 0.613. Lane 9 (HindIII, Pst1) showed bands at Rf 0.20, 0.325, and 0.80. Lane 10 (XhoI, Pst1) showed bands at Rf 0.188, 0.363, and 0.613. Lane 11 (EcoR1, HindIII) showed bands at Rf 0.188, 0.325, and 0.80. Lane 12 (EcoR1, Pst1) showed bands at Rf 0.263, 0.325, and 0.40. The corresponding fragment sizes were used to construct a restriction enzyme map of the plasmid (see Figure 3a). The initial 720-bp nucleotide sequence was searched for restriction enzyme digestion sites using web-based New England BioLabs NEB cutter bioinformatic analysis (see Figure 4).
Following incubation, the Southern blot containing recombinant Bactrocera dorsalis DNA and the control (pUC19 plasmid vector only) were observed. Blue coloration, which indicates positive hybridization of the DIG-UTP probe, was observed in the recombinant DNA blot. No coloration, which indicates no hybridization of the DIG-UTP probe, occurred in the control blot (Figure 5).
The web-based design software Primer3 (http://frodo.wi.mit.edu/) revealed a forward primer of ACTGCCAAAATGTGTGACGA and a reverse primer of TTCTGACCCATACCCACCAT. The segment between primers was 161 base pairs. Following gel electrophoresis of the amplified products of PCR, DNA was observed in lanes 3, 5, 9, and 11 of the 2% agarose gel as bright, high-concentration bands approximately positioned at an Rf-value of 0.77. They can clearly be seen to have similar migration distances, which are all between the 200-bp and 100-bp markers present in lane 7 (standard eGel marker). The negative control samples (containing 0.5 µl of sterile H2O in place of the experimental DNA sequence) in lanes 4, 6, 8, and 10 produced no distinct bands (see Figure 6).
A genome library was created on the plate containing Escherichia coli cells infected with λ-phage containing inserts of genomic Bactrocera dorsalis DNA. A bacterial lawn covered the agar surface, and 45 plaques were observed, as well as two short fingers of no bacterial growth (see Figure 1a).
The plate of cultured E. coli cells containing recombinant DNA plasmids (pUC19 vector with ligated B. dorsalis DNA) produced 476 small circular blue colonies and 512 small circular white colonies. Often the colonies were confluent (Figure 1c).
The agarose gel electrophoresis containing restriction-enzyme-digested DNA samples contained many bands (Figure 2). Lanes 1 and 13 contained 1-kbp DNA marker. Lane 2 (control, pUC19 vector with EcoR1) showed a band at Rf 0.325, lane 3 (BamHI), lane 4 (HindIII), and lane 7 (XhoI) showed bands at Rf 0.125. Lane 5 (Pst1) showed bands at Rf 0.20 and 0.26. Lane 6 (EcoR1) showed bands at Rf 0.175 and 0.325. Lane 8 (EcoR1, XhoI) showed bands at Rf 0.20, 0.325, and 0.613. Lane 9 (HindIII, Pst1) showed bands at Rf 0.20, 0.325, and 0.80. Lane 10 (XhoI, Pst1) showed bands at Rf 0.188, 0.363, and 0.613. Lane 11 (EcoR1, HindIII) showed bands at Rf 0.188, 0.325, and 0.80. Lane 12 (EcoR1, Pst1) showed bands at Rf 0.263, 0.325, and 0.40. The corresponding fragment sizes were used to construct a restriction enzyme map of the plasmid (see Figure 3a). The initial 720-bp nucleotide sequence was searched for restriction enzyme digestion sites using web-based New England BioLabs NEB cutter bioinformatic analysis (see Figure 4).
Following incubation, the Southern blot containing recombinant Bactrocera dorsalis DNA and the control (pUC19 plasmid vector only) were observed. Blue coloration, which indicates positive hybridization of the DIG-UTP probe, was observed in the recombinant DNA blot. No coloration, which indicates no hybridization of the DIG-UTP probe, occurred in the control blot (Figure 5).
The web-based design software Primer3 (http://frodo.wi.mit.edu/) revealed a forward primer of ACTGCCAAAATGTGTGACGA and a reverse primer of TTCTGACCCATACCCACCAT. The segment between primers was 161 base pairs. Following gel electrophoresis of the amplified products of PCR, DNA was observed in lanes 3, 5, 9, and 11 of the 2% agarose gel as bright, high-concentration bands approximately positioned at an Rf-value of 0.77. They can clearly be seen to have similar migration distances, which are all between the 200-bp and 100-bp markers present in lane 7 (standard eGel marker). The negative control samples (containing 0.5 µl of sterile H2O in place of the experimental DNA sequence) in lanes 4, 6, 8, and 10 produced no distinct bands (see Figure 6).
Report Discussion
Discussion:
Our objective was to characterize and sequence an unknown, experimentally obtained segment of genomic DNA from Bactrocera dorsalis. The segment was subjected to several common techniques of genetic analysis (genomic library, amplification of segment in vectors, restriction enzyme digest, bioinformatic analysis of segment, Southern blot to verify inserted sequence, and PCR amplification).
Genomic DNA obtained from the fruit fly Bactrocera dorsalis was treated with EcoRI, which cleaved the genome at each G⇓AATTC recognition site, producing randomly sized, unique DNA fragments. Which were ligated into EMBL3 bacteriophage-λ vector. Once the B. dorsalis fragments were ligated into EMBL3, the recombinant phage was allowed to infect Escherichia coli cells. Once inside the host E. coli cells, the phages replicate by subverting the host’s replication machinery, as phages are not capable of replication outside the host. In this process the bacteriophage DNA may be incorporated into the host cell’s circular DNA strand, as occurs during the lysogenic pathway. If the proper induction event occurs, the cell can be triggered to enter the lytic pathway. During the lytic pathway virus-specific enzymes are created, which allow many replicates of the bacteriophage DNA to be packaged for the infection of other cells. The cell then lyses, and the bacteriophages are released from the cell. When they come in contact with nearby bacterial cells, they land on the cell surface and inject their DNA into the cell cytoplasm. This reinitiates the infection sequence. When many cells are lysed as a result of a series of infections by replicates of one phage particle, a plaque is formed.
The presence of plaques on the bacterial lawn indicates that λ-phage infection was successful. The 45 plaques compose the genome library, and each plaque corresponds to one clonal population of λ-phage with a unique inserted segment of genomic B. dorsalis DNA. One of these plaques was selected for further analysis. The phages present in the plaque were again treated with EcoRI to separate the B. dorsalis DNA from the EMBL3 backbone. The MOI (multiplicity of infection) used in this experiment was far below a 1:1 ratio. If a 1:1 ratio were used, there would be one phage for every one bacterium. All, or nearly all, of the bacteria would die as a result of infection, and no bacterial lawn would be present. The entire agar surface would be plaques of lysed cells, and it would be too difficult to recover one amplified fragment of inserted DNA.
Ligases are enzymes that join the ends of DNA fragments by creating sugar phosphate bonds between the two ends. A vector plasmid (pUC19) containing the lacz gene that produces the enzyme beta-galactosidase and the ampr gene produces enzymes that confer resistance to the antibiotic ampicillin to the host cells. The enzyme beta-galactosidase cleaves X-gal, a synthetic substrate added to the growth medium, and produces products that have a blue color. The restriction enzyme cleaving site (multiple cloning site) is located in the middle of the lacz gene, however. If the foreign DNA fragment is appropriately inserted in this region by way of ligation, the gene will be interrupted and not properly function. Beta-galactosidase will not be produced, and X-gal will not be cleaved to produce a blue color. The colonies will appear white. Also, the presence of colonies confirms the uptake of the plasmid, which contains the ampicillin resistance gene. Without the plasmid, the bacterial cells would not grow on the medium containing ampicillin (LB-amp-Xgal).
476 blue colonies and 512 white colonies were observed on the selective LB-amp-Xgal medium. The presence of colonies on the plate indicated the positive uptake of the plasmids by E. coli cells, although not all of the plasmids contained the recombinant DNA (some were merely pUC19). The white color was indicative of tranformants that contained plasmids with inserted DNA, as the lacz gene was interrupted. The blue color indicated that transformants contained plasmids, but the DNA insert did not interrupt the lacz gene (as these cells still produced beta-galactosidase).
The most likely explanation for the continued production of beta-galactosidase is that B. dorsalis DNA was not inserted into the plasmid, but it is also possible that the DNA was ligated into another region of the plasmid vector (unlikely). A blue colony could occur if the insert DNA adjacent to the longer sequence of the interrupted beta-galactosidase gene was sufficiently similar to the sequence that had been there before cleavage and ligation. The gene might still properly function, cleaving X-gal into blue products. It is also possible that the insert DNA could contain the beta-galactosidase gene.
Restriction enzymes, which cleave DNA at specific sequences (recognition sites), can help characterize unknown DNA segments. By treating a sample of DNA with a combination of single and double digests, a restriction enzyme map can be created, indicating the approximate position of known recognition sites along the target sequence.
Restriction enzymes were selected that did not have recognition sites in the pUC19 vector (other than MCS) in order to facilitate map creation. Bam HI, EcoR1, HindIII, and Pst1 had recognition sites in the MCS of pUC19. Xho1 did not have any recognition site in the pUC19 vector.
After electrophoresing the DNA fragment products of restriction enzyme digests, the sizes of each fragment were calculated by comparison with the 1kb marker. A restriction enzyme map of the recombinant plasmid was constructed. The pUC19 backbone is approximately 2700 bp in size. The recombinant plasmid had a size of 8700 bp as indicated by the fragment sizes of BamHI and HindIII digests, which both cleaved the circular plasmid only once. The insert DNA was around 6000 bp. The HindIII restriction site was located around 500-bp from the MCS, the Xho1 restriction site was around 1000 bp from the MCS, and the Pst1 restriction site was 3550 bp from the MCS. EcoR1 restriction sites occurred at both interfaces of the pUC19 vector and the B. dorsalis DNA insert. This constructed restriction enzyme map was compared to a computer-generated map (Figure 4) for 720-bp of the insert sequence. A HindIII restriction site was found at bp 175 of the 720-bp sequence. This indicates that the 720-bp sequence did not originate at the MCS, but approximately 545 bp distal to it. The Xho1 site was not seen in the 720-bp sequence.
Probes and Southern blots are useful tools in their ability to verify the presence of a specific DNA sequence. A probe can be constructed that matches the target sequence. A template strand that matches the target sequence is amplified using PCR. Initially, denatured sample DNA is dotted onto a nylon membrane and crosslinked using UV radiation. Next, prehybridization is performed by wetting the membrane and blocking sites that could allow non-specific binding by the probe. Following prehybridization, the denatured probe DNA is added to the membrane-bound samples. Hybridization occurs between the probe and its complimentary target sequence. Following hybridization, the samples are washed several times in order to remove non-bound and weakly bound probe sequences. Following the wash sequences, the specific binding of the probe to the target sequence can be detected.
The mechanism of detection of DNA sequences using the Southern blot method is simple, although there are several steps. Initially, a probe is created that contains the nucleotide sequence under investigation. During hybridization, this probe sequence will bind complementarily to the target sequence if it is present in the sample. A “tag” or label is necessarily built into the probe to verify hybridization with the target sequence. One type of tag involves the use of DIG-UTP probe, in which a modified nucleotide (DIG-UTP) is used to synthesize the probe during PCR. Once the probe hybridizes to the target sequence on the membrane, the sample is treated with an enzyme-bound (alkaline phosphatase) antibody that is directed against the DIG modification. The antibody and the enzyme adhere to the DIG-UTP nucleotides. When enzyme-specific substrates are added to the sample, the presence of the target sequence can be verified by formation of the products, which often designed to yield a change in color (in this case blue).
A dot blot was conducted on the experimental recombinant sample in order to verify the presence of the actin-coding gene in Bactrocera dorsalis. After ligation and cloning of the insert in the pUC19 vector, verification of the target sequence is necessary to validate the results obtained. The results indicated that a blue coloration was produced in the sample containing recombinant DNA. The presence of the target sequence was verified, by cleavage of the added substrates and the formation of blue products. This cleavage was performed by the enzyme conjugated to the antibody, which was directed toward (and adhered to) the DIG-UTP nucleotides present in the previously synthesized probe.
The negative control sample did not produce any blue coloration. This indicates that cleavage of the substrates did not occur, and therefore, the enzyme responsible for cleavage was not present. All of the enzyme-conjugated antibodies were washed off, as there were no DIG-UTP nucleotides present in the sample. The probe did not bind to any sequences in the sample, and the target sequence was not detected in the pUC19 vector. It can also be concluded that no non-specific binding occurred by the probe or the antibody.
Polymerase chain reaction (PCR) amplification is a convenient method of amplifying a section of DNA in vitro. In contrast to in vivo cloning of a plasmid containing a ligated DNA insert, PCR can produce billions of copies of the target DNA sequence in a very short time with little effort required for isolation of the sequence after amplification. The standard cycle of PCR amplification involves the repetition of three steps: denaturation (during which the DNA strand is denatured to produce two template strands), annealing (during which pre-selected primers bind to the target sequences), and extension (during which the DNA polymerase elongates the DNA strand through complimentary base pairing of the dNTPs provided in the reaction mixture). The key to the PCR process is the use of a heat-stable DNA polymerase. Taq polymerase, isolated from Thermus aquaticus, is heat-stable, and therefore survives the denaturation stage of the cycle.
The presence of clearly visible, high-intensity bands in the lanes containing the experimental DNA (lanes 3, 5, 9, and 11) indicates that some sequence of DNA was amplified by the PCR process. It also indicates that the primers, polymerase, and dNTPs functioned properly and the buffer and MgCl2 concentration were suitable for amplification. Based on the migration distance of the band, which was greater than the 200-bp marker and less than the 100-bp marker, it can be concluded that the amplified sequence was between 100 and 200 base pairs in length. This corresponds to our hypothesized amplified segment length of 161 base pairs (this length was determined by the selection and design of primers 1 and 2). The results were positive, indicating the target sequence was present and sufficiently amplified.
The electrophoresed negative control samples that contained only the PCR amplification mixture and 0.5 µl of H2O in place of DNA (lanes 4, 6, 8, and 10) produced no distinct bands. This indicates that no sequence of DNA was amplified during the PCR process. If the sample had been contaminated by foreign DNA, this DNA would have possibly been amplified (if it contained a sequence similar to that of primers 1 and/or 2 or if those primers bound non-specifically) as indicated by the presence of a band in the electrophoresis gel. The negative control produced no results, indicating no significant contamination.
In conclusion, these techniques were combined to characterize and confirm the unknown sequence of B. dorsalis DNA as a gene that codes for the protein actin, a highly conserved protein among species that functions in the cytoskeleton and in muscular contraction of B. dorsalis cells.
Our objective was to characterize and sequence an unknown, experimentally obtained segment of genomic DNA from Bactrocera dorsalis. The segment was subjected to several common techniques of genetic analysis (genomic library, amplification of segment in vectors, restriction enzyme digest, bioinformatic analysis of segment, Southern blot to verify inserted sequence, and PCR amplification).
Genomic DNA obtained from the fruit fly Bactrocera dorsalis was treated with EcoRI, which cleaved the genome at each G⇓AATTC recognition site, producing randomly sized, unique DNA fragments. Which were ligated into EMBL3 bacteriophage-λ vector. Once the B. dorsalis fragments were ligated into EMBL3, the recombinant phage was allowed to infect Escherichia coli cells. Once inside the host E. coli cells, the phages replicate by subverting the host’s replication machinery, as phages are not capable of replication outside the host. In this process the bacteriophage DNA may be incorporated into the host cell’s circular DNA strand, as occurs during the lysogenic pathway. If the proper induction event occurs, the cell can be triggered to enter the lytic pathway. During the lytic pathway virus-specific enzymes are created, which allow many replicates of the bacteriophage DNA to be packaged for the infection of other cells. The cell then lyses, and the bacteriophages are released from the cell. When they come in contact with nearby bacterial cells, they land on the cell surface and inject their DNA into the cell cytoplasm. This reinitiates the infection sequence. When many cells are lysed as a result of a series of infections by replicates of one phage particle, a plaque is formed.
The presence of plaques on the bacterial lawn indicates that λ-phage infection was successful. The 45 plaques compose the genome library, and each plaque corresponds to one clonal population of λ-phage with a unique inserted segment of genomic B. dorsalis DNA. One of these plaques was selected for further analysis. The phages present in the plaque were again treated with EcoRI to separate the B. dorsalis DNA from the EMBL3 backbone. The MOI (multiplicity of infection) used in this experiment was far below a 1:1 ratio. If a 1:1 ratio were used, there would be one phage for every one bacterium. All, or nearly all, of the bacteria would die as a result of infection, and no bacterial lawn would be present. The entire agar surface would be plaques of lysed cells, and it would be too difficult to recover one amplified fragment of inserted DNA.
Ligases are enzymes that join the ends of DNA fragments by creating sugar phosphate bonds between the two ends. A vector plasmid (pUC19) containing the lacz gene that produces the enzyme beta-galactosidase and the ampr gene produces enzymes that confer resistance to the antibiotic ampicillin to the host cells. The enzyme beta-galactosidase cleaves X-gal, a synthetic substrate added to the growth medium, and produces products that have a blue color. The restriction enzyme cleaving site (multiple cloning site) is located in the middle of the lacz gene, however. If the foreign DNA fragment is appropriately inserted in this region by way of ligation, the gene will be interrupted and not properly function. Beta-galactosidase will not be produced, and X-gal will not be cleaved to produce a blue color. The colonies will appear white. Also, the presence of colonies confirms the uptake of the plasmid, which contains the ampicillin resistance gene. Without the plasmid, the bacterial cells would not grow on the medium containing ampicillin (LB-amp-Xgal).
476 blue colonies and 512 white colonies were observed on the selective LB-amp-Xgal medium. The presence of colonies on the plate indicated the positive uptake of the plasmids by E. coli cells, although not all of the plasmids contained the recombinant DNA (some were merely pUC19). The white color was indicative of tranformants that contained plasmids with inserted DNA, as the lacz gene was interrupted. The blue color indicated that transformants contained plasmids, but the DNA insert did not interrupt the lacz gene (as these cells still produced beta-galactosidase).
The most likely explanation for the continued production of beta-galactosidase is that B. dorsalis DNA was not inserted into the plasmid, but it is also possible that the DNA was ligated into another region of the plasmid vector (unlikely). A blue colony could occur if the insert DNA adjacent to the longer sequence of the interrupted beta-galactosidase gene was sufficiently similar to the sequence that had been there before cleavage and ligation. The gene might still properly function, cleaving X-gal into blue products. It is also possible that the insert DNA could contain the beta-galactosidase gene.
Restriction enzymes, which cleave DNA at specific sequences (recognition sites), can help characterize unknown DNA segments. By treating a sample of DNA with a combination of single and double digests, a restriction enzyme map can be created, indicating the approximate position of known recognition sites along the target sequence.
Restriction enzymes were selected that did not have recognition sites in the pUC19 vector (other than MCS) in order to facilitate map creation. Bam HI, EcoR1, HindIII, and Pst1 had recognition sites in the MCS of pUC19. Xho1 did not have any recognition site in the pUC19 vector.
After electrophoresing the DNA fragment products of restriction enzyme digests, the sizes of each fragment were calculated by comparison with the 1kb marker. A restriction enzyme map of the recombinant plasmid was constructed. The pUC19 backbone is approximately 2700 bp in size. The recombinant plasmid had a size of 8700 bp as indicated by the fragment sizes of BamHI and HindIII digests, which both cleaved the circular plasmid only once. The insert DNA was around 6000 bp. The HindIII restriction site was located around 500-bp from the MCS, the Xho1 restriction site was around 1000 bp from the MCS, and the Pst1 restriction site was 3550 bp from the MCS. EcoR1 restriction sites occurred at both interfaces of the pUC19 vector and the B. dorsalis DNA insert. This constructed restriction enzyme map was compared to a computer-generated map (Figure 4) for 720-bp of the insert sequence. A HindIII restriction site was found at bp 175 of the 720-bp sequence. This indicates that the 720-bp sequence did not originate at the MCS, but approximately 545 bp distal to it. The Xho1 site was not seen in the 720-bp sequence.
Probes and Southern blots are useful tools in their ability to verify the presence of a specific DNA sequence. A probe can be constructed that matches the target sequence. A template strand that matches the target sequence is amplified using PCR. Initially, denatured sample DNA is dotted onto a nylon membrane and crosslinked using UV radiation. Next, prehybridization is performed by wetting the membrane and blocking sites that could allow non-specific binding by the probe. Following prehybridization, the denatured probe DNA is added to the membrane-bound samples. Hybridization occurs between the probe and its complimentary target sequence. Following hybridization, the samples are washed several times in order to remove non-bound and weakly bound probe sequences. Following the wash sequences, the specific binding of the probe to the target sequence can be detected.
The mechanism of detection of DNA sequences using the Southern blot method is simple, although there are several steps. Initially, a probe is created that contains the nucleotide sequence under investigation. During hybridization, this probe sequence will bind complementarily to the target sequence if it is present in the sample. A “tag” or label is necessarily built into the probe to verify hybridization with the target sequence. One type of tag involves the use of DIG-UTP probe, in which a modified nucleotide (DIG-UTP) is used to synthesize the probe during PCR. Once the probe hybridizes to the target sequence on the membrane, the sample is treated with an enzyme-bound (alkaline phosphatase) antibody that is directed against the DIG modification. The antibody and the enzyme adhere to the DIG-UTP nucleotides. When enzyme-specific substrates are added to the sample, the presence of the target sequence can be verified by formation of the products, which often designed to yield a change in color (in this case blue).
A dot blot was conducted on the experimental recombinant sample in order to verify the presence of the actin-coding gene in Bactrocera dorsalis. After ligation and cloning of the insert in the pUC19 vector, verification of the target sequence is necessary to validate the results obtained. The results indicated that a blue coloration was produced in the sample containing recombinant DNA. The presence of the target sequence was verified, by cleavage of the added substrates and the formation of blue products. This cleavage was performed by the enzyme conjugated to the antibody, which was directed toward (and adhered to) the DIG-UTP nucleotides present in the previously synthesized probe.
The negative control sample did not produce any blue coloration. This indicates that cleavage of the substrates did not occur, and therefore, the enzyme responsible for cleavage was not present. All of the enzyme-conjugated antibodies were washed off, as there were no DIG-UTP nucleotides present in the sample. The probe did not bind to any sequences in the sample, and the target sequence was not detected in the pUC19 vector. It can also be concluded that no non-specific binding occurred by the probe or the antibody.
Polymerase chain reaction (PCR) amplification is a convenient method of amplifying a section of DNA in vitro. In contrast to in vivo cloning of a plasmid containing a ligated DNA insert, PCR can produce billions of copies of the target DNA sequence in a very short time with little effort required for isolation of the sequence after amplification. The standard cycle of PCR amplification involves the repetition of three steps: denaturation (during which the DNA strand is denatured to produce two template strands), annealing (during which pre-selected primers bind to the target sequences), and extension (during which the DNA polymerase elongates the DNA strand through complimentary base pairing of the dNTPs provided in the reaction mixture). The key to the PCR process is the use of a heat-stable DNA polymerase. Taq polymerase, isolated from Thermus aquaticus, is heat-stable, and therefore survives the denaturation stage of the cycle.
The presence of clearly visible, high-intensity bands in the lanes containing the experimental DNA (lanes 3, 5, 9, and 11) indicates that some sequence of DNA was amplified by the PCR process. It also indicates that the primers, polymerase, and dNTPs functioned properly and the buffer and MgCl2 concentration were suitable for amplification. Based on the migration distance of the band, which was greater than the 200-bp marker and less than the 100-bp marker, it can be concluded that the amplified sequence was between 100 and 200 base pairs in length. This corresponds to our hypothesized amplified segment length of 161 base pairs (this length was determined by the selection and design of primers 1 and 2). The results were positive, indicating the target sequence was present and sufficiently amplified.
The electrophoresed negative control samples that contained only the PCR amplification mixture and 0.5 µl of H2O in place of DNA (lanes 4, 6, 8, and 10) produced no distinct bands. This indicates that no sequence of DNA was amplified during the PCR process. If the sample had been contaminated by foreign DNA, this DNA would have possibly been amplified (if it contained a sequence similar to that of primers 1 and/or 2 or if those primers bound non-specifically) as indicated by the presence of a band in the electrophoresis gel. The negative control produced no results, indicating no significant contamination.
In conclusion, these techniques were combined to characterize and confirm the unknown sequence of B. dorsalis DNA as a gene that codes for the protein actin, a highly conserved protein among species that functions in the cytoskeleton and in muscular contraction of B. dorsalis cells.
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