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Genomic editing by subcloning of the cas9 gene
- 2. Abstract
The CRISPR/Cas9 system is part of the adaptive immune response of certain
prokaryotes, protecting the organism by sitespecific cleavage of invading DNA. The Cas9 gene
is currently being researched for use as a genome editing tool, targeting specific genomic
sequences. The Cas9 gene can be injected with snRNA into cells for genome editing, to either
knockout or replace a gene. In a series of exercises, the Cas9 gene was subcloned from the
pCAMBIACas9 snRNA plasmid into pcDNA 3.1/V5His C. After recombination of the
pcDNA3.1/HisCCas9 using onestep annealing Gibson cloning, the plasmid was transformed
into competent E. Coli cells. The new expression vector was used for invitro transcription of
the Cas9 gene. The objective of this experiment was to subclone a codon optimized CRISPR
associated protein 9 (Cas9) gene from Streptococcus pyogenes into the expression plasmid
pcDNA3.1/V5HisC. The length of the pcDNA3.1/V5HisC restriction digest and Cas9 PCR
product were calculated using regression analysis. The length of the pcDNA3.1/V5HisC
restriction digest was calculated to be 6471 base pairs in length, and the Cas9 PCR was
calculated to be 5446 base pairs in length. The DNA purification resulted in recovery of 20 ng
of Cas9 DNA at an A260/A280 ratio of 3.23. and 5ng of pcDNA3.1/V5HisC restriction digest at an
A260/A280 ratio of 0.18. Transformation of the two positive control competent cells had
transformation efficiencies of 200 and 320, for the plates with a dilution factor of 20 and 5
respectively. The absolute amounts of recombinant pcDNA3.1/HisCCas9 recombinant plasmid
was 26.85 ng and the A260/A280 ratio equaled 1.04. The length of the plasmid was calculated to be
8256 base pairs. The dot blot confirmed the recombination presence of the recombinant
pcDNA3.1/HisCCas9 plasmid in four experimental samples.
Introduction
An adaptive immune response in prokaryotes, used to combat foreign DNA, has become
of particular interest because of the ability to manipulate the system for genomic editing. The
response includes clustered regulatory interspaced short palindromic repeats (CRISPR) and
CRISPRassociated genes (Cas genes), which protect the cell against invading viruses and
plasmids. This system is of interest for genome editing, gene knockout and replacement in
1
- 3. particular, because it entails targeting specific genomic sequences and cleaving at designated
sites on the sequence.
There are three main types of CRISPR/Cas systems, with type II being the most
extensively studied. The type II system contains the Cas9 gene, which works in conjunction
with CRISPRRNA (crRNA) and trans activating RNA (tracerRNA) to cleave foreign DNA.
The immune response must be preceded by cleaving and incorporating small pieces of the DNA
between the CRISPR locus, which are a series of short base pair repeats located on a plasmid in
the organism. When invaded by the same virus or plasmid, the organisms CRIPSRCas system
responds. The first part of the process transcribes short CRIPSR RNAs (crRNAs), which contain
genetic information complementary to the invader (1), and transactivating crRNA (tracer RNA)
containing a short sequence complementary to crRNA. In turn the Cas gene, located near the
CRISPR sequence, is transcribed forming the Cas9 endonuclease. The Cas9, crRNA, and tracer
RNA form a complex, and the crRNA is used to locate the invading virus or plasmid and cleave
the genetic material of the invader at a particular site (1). One important aspect of the system is
that it targets only foreign DNA, distinguishing between self and nonself. If indiscriminate, the
CRISPR locus on the organism's DNA would be cleaved, but this is avoided because the Cas9
gene cleaves at a protospacer adjacent motif (PAM) site. One important aspect of the PAM site
is that it allows for double stranded DNA cleavage (1), important for cleavage of plasmids and
double stranded DNA viruses.
The CRISPRCas9 system has implications beyond being an adaptive immune response
in prokaryotes; in particular it can be utilized for genomic editing. The Cas9 system is being
engineered to knockout genes by creating a sitespecific double stranded break (2). Using an
engineered CRISPR/Cas9 system, a double stranded break of target DNA can be accomplished
(2), and this double stranded break initiates nonhomologous end joining repair resulting in either
deletions or insertions. By causing these changes in the target gene, the gene is then no longer
functional. In addition it can be manipulated to repair genes, using the CRISPR/Cas9 system to
cause nicks in the target gene (2). In addition to the CRISPR/Cas9 complex, a donor template of
DNA containing homologous ends to the nicks is inserted with the template containing genomic
2
- 4. information for insertion into the genome (2). These nicks are repaired using homologous end
joining repair, and the donor template is incorporated into the genome during the repair process.
This form of genomic editing is proving to be a powerful tool. However, in order to
utilize and further study the abilities of the CRISPR/Cas9 system, the genome must be isolated
and reproduced in large quantities, which is the focus of the experiments that were performed.
The goal of the experiments was to subclone the Cas9 gene, located in the pCAMBIACas9
snRNA plasmid of Streptococcus pyogenes, into pcDNA3.1/V5His C, creating the new
expression vector pcDNA3.1/HisCCas 9. The new recombinant plasmid was then used to
transform E. coli, in order to express the Cas9 gene. This will allow for utilization of Cas9, in
combination with snRNA with target sequences, for genomic editing purposes.
In order to accomplish this, the pcDNA 3.1 plasmid was digested with a BAMHI
restriction enzyme in order to linearize the plasmid. In conjunction, the Cas9 gene on the
pCAMBIACas9 plasmid DNA was amplified using Polymerase Chain Reaction (PCR). In
order to confirm that the plasmid was digested and the PCR amplification of the Cas9 gene was
successful, an agarose gel was run, comparing the size of the molecular weight marker to the
weights of the digest (5498 base pairs in size), and Cas9 gene (4194 base pairs in size). The
PCR amplifications of the Cas9 gene were then purified to ensure that only the Cas9 gene, and
no other byproducts, were present.
After purification, the Cas9 gene was again amplified, this time utilizing Gibson Cloning.
Gibson cloning allowed for recombination of the Cas9 gene by utilization of homologous
recombination. The Cas9 gene was amplified with specific primers that overlap the BAM HI site
of the linearized pcDNA 3.1 plasmid. Taq DNA polymerase and T4 DNA ligase combined the
Cas9 gene with the linearized pcDNA 3.1/V5HisC plasmid to create the pcDNA3.1/HisCCas9
plasmid (9.6 kilobases in size). The new plasmid was transformed into competent E. coli cells
by mixing the cells and the plasma and briefly heat shocking the cells.
Once the cells had been transformed, it was necessary to confirm the quality and quantity
of the recombinant pcDNA3.1/HisCCas9 plasmid. The quality was determined by extracting
and linearizing the recombinant pcDNA3.1/HisCCas9 plasmid DNA, using restriction enzymes,
and running an agarose gel to confirm the size of the DNA. The size of linearized DNA and the
3
- 5. parent pcDNA3.1/V5HisC plasmid was calculated using regression line, generated using the
molecular weight marker. The quantity of the plasmid DNA was done through the use of UV
spectroscopy, which allowed for calculation of the concentration of the DNA and the
concentration was used to calculate the quantity of DNA. The final step in confirming that the
plasmid created was the recombinant pcDNA3.1/HisCas9 plasmid DNA was using a Southern
dot blot. In the dot plot a small amount of the denatured recombinant plasmid DNA, in addition
to a positive and negative control, was spotted onto a nylon membrane and hybridized with a
probe containing a complementary base pair sequence. Visualization, using a color substrate
solution, was used to confirm the presence of the recombinant pcDNA3.1/HisCas9 plasmid.
Methods
PCR Synthesis
The first process in this experiment was PCR synthesis of the Cas9 sequence. PCR
consists of the production or amplification of billions of copies of the target gene sequence. PCR
reaction includes DNA polymerase, DNA template, deoxyribonucleotides, primers to flank the
sequence of interest, and a buffer containing essential salts and ions. Denaturation, annealing and
extension are repeated multiple times throughout PCR.
Initial setup involved thawing the DNA template, 2X High Fidelity Phusion Buffer and
primers. Thawing was required to allow for mixing, these components were initially frozen to
maintain their integrity before using them in the lab. These components were mixed and
centrifuged, ensuring homogeneity. It was necessary to assemble a PCR Master Mix to amplify
the Cas9 gene from the plasmid by using a final concentration of 43.75 µl 2X High Fidelity
Phusion Buffer, 19.25 µl nucleasefree dH₂O, 4.37 µl 10µM Cas9 forward and reverse primer
mix, 2.625 µl DMSO for a final total volume of 70 µl in a 0.5 mL microfuge tube. It was
necessary to add the phusion buffer last to ensure that the exonuclease in the buffer would not
degrade the chosen primers. The Master Mix included phusion buffer containing phusion
polymerase to replicate DNA and check for errors in the genomic sequence, while dH₂O was in
the master mix to ensure that the catalytic activity of polymerases chose the specific targeted
DNA. Primers were included in the Master Mix to amplify a certain area of the Cas9 and code
4
- 6. for both the 5’ and the 3’ end of the DNA strands. DMSO was used in the Master mix as an
enhancing agent in PCR reactions due to its ability to separate DNA strands.
Then, 5µL of Cas9 template DNA, consisting of pCambiaCas9 and sgRNA plasmid,
were pipetted into three separate 0.2 mL microfuge tubes along with 20µL of Master Mix. Three
separate PCR reactions were performed to increase the chance of amplifying the Cas9 gene and
obtaining a pure product from the PCR reaction. Those three PCR reactions were placed into the
thermocycler and programmed to perform in this way: Initial denaturation at 98°C for 30 seconds
allowing denaturation of the DNA strands by breaking the hydrogen bonds and opening up the
strands, denaturation at 98°C for 10 seconds, primer annealing at 55°C for 30 seconds allowing
primers to anneal to specific regions of DNA, and extension at 72°C for 2 minutes allowing the
phusion polymerase to extend the primer by adding nucleotides to the developing DNA strand.
These cycles of denaturation, annealing and extension were ran thirty times, producing a high
yield of the selected Cas9 sequence. One final extension was ran at 72°C for 10 minutes to allow
any partial sequences that remained to complete their sequences. The sample was placed in an
ice bath at 4°C to be used later.
After those initial steps were completed, it was necessary to assemble a single enzyme
digest to be kept on ice: 7µL of H2O, 2µL of 10X Reaction Buffer, 10µL of pcDNA3.1/V5His
C (0.30 ng/µL), and 1 µL of BAM HI for a total of 20.0µL. In two 0.5mL microfuge tubes, 300
ng of pcDNA3.1/V5His C was digested in 20 uL volume using the restriction enzyme BAM HI.
The BAM HI is used to linearize the expression vector a specific site, regions of overhang in
BAM HI match up with the 5’ ends of the Cas9 primers. The components were added to the
microfuge tube in the order listed previously. The reactions were centrifuged briefly to combine
components and placed in a 37°C water bath and incubated for an hour to allow optimal
activation of BAM HI.
Agarose Gel Electrophoresis
Once PCR synthesis was completed, agarose gel electrophoresis was started. The agarose
gel helped to check whether the PCR synthesis of Cas9 made any product. In recombinant DNA,
it is necessary to separate DNA molecules from one another and from contaminating substances,
5
- 7. which can be done by gel electrophoresis. The DNA molecules are separated by fragment size,
which can be compared with fragments of a known size.
This began by assembling the electrophoresis unit with a comb in the casting bed and
making the agarose gel. The 1% agarose gel was made with 0.50 g of agarose and 50 mL of 1X
TAE buffer in a 125 mL Erlenmeyer flask. The 1X TAE buffer was made from a 50X TAE stock
composed of 242 g Trisbase, 57.1 mL glacial acetic acid, 100 mL 0.5 M EDTA pH 8.0, and 1
mL H20 using a 1:50 dilution, totaling 1 mL 50X TAE stock and 49 mL of H₂O. This mixture
was placed into the microwave at full power until it just began to boil. Before it boiled over, it
was taken out and mixed gently by swirling then placed back in the microwave. This process was
repeated until the solution boiled and had no “lenses”, meaning no visible particles in the clear
solution. Once no “lenses” were visible, the agarose gel had completely dissolved and set on the
table top until slightly cooled. Once the solution was cool enough to touch, the gel was poured
into the gel bed in the electrophoresis box. The gel stood close to 30 minutes until it was firm
and translucent. The casting bed was repositioned in the gel box and 1X TAE Buffer was added
to each side of the box, enough buffer was added to slightly cover the gel. The comb was
carefully pulled out of the gel to create the wells.
To the agarose gel, 12 µL of DI water was added to the tube containing 3 µL of
Lambda/Hind III size marker (100ng/µL), while 3 µL of 6X DNA loading dye (30% glycerol,
0.25% Bromophenol Blue, 0.25% Xylene cyanol, 6X SYBR green) was added to the marker and
mixed. This was used as the marker to compare molecular weights with the actual results. The
loading dye bonds to the nucleic acid and the electrophoresis separates the solution by molecular
weight, creating a visual representation of whether the PCR synthesis made any product.
The Cas9 PCR reactions were combined into a single tube, 15 µL of this combined PCR
reaction was loaded into one of the wells to check the PCR reaction product. The remaining PCR
was frozen in 20°C to be used in the next part of the experiment. At least 3 µL of loading dye
was added to each sample to bring the final concentration to 1X. The two pcDNA3.1/V5His C
restriction digest reactions were combined into one tube. 10uL was moved into one tube along
with 5 µL of water and 3 µL of 6X DNA loading dye to be used in the gel electrophoresis. The
remainder of the sample was frozen in 20°C freezer to be used in the next part of the
6
- 8. experiment. Using the P20 pipette, 18µL from each tube was placed into a well in the agarose
gel. The top was placed onto the electrophoresis box and the leads connected appropriately, red
to red and black to black. The power was turned on and the gel ran at 5V/cm for one hour. The
gel was taken out at the end of the electrophoresis run and examined using a transilluminator.
The transilluminator uses a UVB light source to allow the DNA with the dye to light up and
become visible.
QIAQuick DNA Extraction
While the gel electrophoresis was running, the QIAQUICK DNA purification from
PCR/Restriction digest was completed. QIAQUICK DNA purification is used to purify PCR
amplification and DNA digestions products. It was necessary to add ethanol to the Buffer PE
before use to cause the DNA to join with positive ions and form a precipitate. 5 volumes of
Buffer PB were added to 1 volume of each tube containing the PCR reaction and restriction
digest. Buffer PB allows for efficient binding of PCR products to the spincolumn. The color of
our sample was good, but it would have been necessary to add 10µL of 3M sodium acetate (pH
5.0) and mix to create the correct color of yellow. A QIAquick spin column was placed in a 2mL
collection tube. To bind the DNA, the sample was added to the QIAquick column and
centrifuged for 1 min, flowthrough was discarded, and centrifuged once more at 13,000 RPM to
remove the residual wash buffer. The QIAquick column was placed into a clean microcentrifuge
tube. To elute the DNA, 50 uL of Buffer EB (10 mM trisCl, pH 8.5) was added to the center of
the QIAquick membrane, allowed to stand for 4 minutes and then centrifuged for 1 minute. The
amount of DNA was quantitated using a nanodrop. A nanodrop is a spectrophotometer that is
used to quantify and assess the purity of DNA.
Gibson Cloning
The next step was Gibson Cloning. Gibson cloning is a way to efficiently connect two
pieces of DNA with overlapping ends regardless of fragment length or end compatibility. Gibson
cloning works by joining multiple DNA fragments in a single isothermal reaction First an
exonuclease cleaves nucleotides at the 5’ end, annealing DNA fragments from 3’ to 5’, DNA
polymerase comes along and closes gaps in the annealed fragment and then DNA ligase joins the
7
- 10. with a negative control and no DNA, and one with the experimental Gibson Cloning reaction and
pcDNA3.1/V5His CCas9. It is necessary to use three different transformation reactions, the
negative control tested the efficacy of ampicillin, the positive control tested for competency of
cells, and the experimental plate tested for successful transformation or recombination. Each of
the tubes were gently stirred with the pipet tip and incubated on ice for two minutes. The tubes
were transferred to a 42°C water bath for 30 seconds and placed on ice for 5 minutes, then 950
uL of SOC medium was added.
Heat shock facilitates transformation along with SOC medium. 50 uL and 200 uL of each
suspension was added to each of two LB plates supplemented with 100 ug/mL ampicillin and
spread gently with sterile glass balls. Some plasmids are ampicillin resistant, therefore DNA with
an ampicillin resistance will be able to grow on the plates and those without the ampicillin
resistance will not be able to grow. The plates were incubated at 37°C overnight to allow
formation of colonies. Once the plates had been incubated, several colonies were subcultured in
2.5 mL of ampicillin supplemented (100 ug/mL) LB culture tubes. Subculturing was used to
expand the number of cells in the culture.
Following the bacterial transformation was the plasmid miniprep. The QIAprep miniprep
procedure produces a sufficiently pure plasmid to allow endonuclease digestion. Four phase
cultures of E.Coli were grown from recombinant colonies, 1.5 mL of each was transferred to a
microfuge tube and centrifuged at 15000g for 30 seconds and the supernatants were discarded.
Then, 5 mL of bacterial overnight culture was pelleted by centrifugation at >8000 rpm for 3
minutes at room temperature. Pellets are an isolated specimen that allows further analyzation.
The pelleted bacterial cells were completely resuspended in 250 µL Buffer P1 and transferred to
a microcentrifuge tube, afterwards making sure that no clumps of cells remained. Buffer P1 is
used to purify plasmid DNA.
Then, 250 µL of Buffer P2 was added and mixed by inverting the tube about 12 times
until the solution turned blue. Buffer P2 is used a lysis buffer while preparing plasmid DNA.
This was incubated for 5 minutes, but no longer than 5 minutes to speed up the lysis process. The
high salt buffers in the lysis procedures make sure that only DNA binds to the membrane while
RNA, cellular proteins, and metabolites flowthrough the membrane. 350 µL of Buffer N3 was
9
- 11. added and mixed immediately by inverting about 12 times until the solution turned colorless.
This was centrifuged for 10 minutes. This supernatant was applied to the QIAprep spin column
by pipetting and centrifuged for 30 seconds, discarding the flowthrough. The QIAprep spin
column was washed and centrifuged twice, once with 500 uL Buffer PB and once with 750 µL
Buffer PE. Then, the QIAprep spin column was transferred to the collection tube and centrifuged
for 1 minute to reduce residual wash buffer. Finally, the QIAprep spin column was placed into a
clean 1.5 mL microcentrifuge tube and the DNA was eluted by adding 50 uL Buffer EB (10 MM
TrisCl, pH 8.5) to the center of the spin column, allowed to stand for 1 min, and centrifuged for
1 min. The series of wash steps efficiently removes endonucleases and salts before high quality
plasmid DNA is eluted from the membrane.
UV Spectroscopic Analysis
Ultraviolet spectroscopic analysis of nucleic acids was used to measure the DNA sample
at different wavelengths to assess the concentration and purity of the nucleic acid. Ultraviolet
spectroscopes use UV light absorption to measure the attenuation of a beam after it passes
through a sample. The absorbance measurement of nucleic acids is 260 nm as long as
contributions from contaminants and buffer components are taken into consideration. Although
absorbance readings cannot tell the difference between DNA and RNA, the ratio of absorbance
values at 260 and 280 nm can be used to indicate purity. The first step was preparing 200 µL of a
1:40 dilution of dissolved DNA by adding 5 µL of concentrated solution to 195 µL of dH₂O in a
microfuge tube. This was mixed by vortexing.
The absorbance was measured with the spectrophotometer, using water as the blank. The
absorbance was measured at both 260 nm and 280 nm. If the A260 value was <0.1 or >1.0,
appropriate dilutions would need to be made so the reading falls into that range. The DNA
concentration and A260/A280 provides an estimate of the concentration and purity of the DNA
sample, as mentioned previously. An A260 value of 1.0 represents a pathlength of 1cm and a
concentration of DNA in solution of 50 µg/mL. Pure DNA typically has a A260/A280 ratio between
1.7 and 1.9 dependent upon the base composition.
10
- 13. making a 0.5 cm x 0.5 cm grid using a pencil. Approximately 20 mL of 6X SSC was poured into
a petri dish and the membrane placed on the surface, allowing it to submerge. This process is
used to allow DNA to accumulate on the surface of the membrane, allowing accessibility to the
probe and in higher concentrations than in an agarose gel. The membrane was incubated for 10
minutes to ensure that the membrane was fully saturated.
For each plasmid sample, positive and negative controls, 200 µL of a spotting solution
containing 100 ng/µL of DNA needed to be made to show up on the membrane. Volumes were
calculated by using the equation C1V1= C2V2. The final concentration to made for our experiment
was 5 µL of 20X SSC, 4.1 µL of plasmid DNA, 1 µL of 1 M NaOH, 1.25 µL of 200 mM EDTA,
and 13.65 µL of diH₂O for a total volume of 25 µL. The sample was heated at 100°C for 10
minutes, then placed on ice. The high temperature and spotting solution denatured the DNA into
single strands. The wetted membrane was placed onto clean paper towels to dry. Each sample
was spun in the microcentrifuge for 5 seconds, and 100 ng of each sample spotted onto the
membrane using a pipet and allowed to dry. 2 µL was spotted in each application and applied
twice, allowing each spot to dry before reapplying. The membrane was rinsed briefly in 2X SSC
and allowed to air dry before UVcross linking the DNA. The 2X SSC is used to transfer the
DNA to the surface of the membrane. The membrane was dried in a dark drawer at room
temperature.
Hybridization and washing consisted of placing the membrane and 20 mL standard
hybridization solution into a SealAMeal bag. The bag was placed into a hybridization oven for
3 hours at 52℃. The hybridization was completed by Dr. Read, these are assumptions on the
way that Dr. Read proceeded with this portion of the process including the amount of solution,
time, and temperature. The probe was denatured in a boiling water bath for five minutes, and was
subsequently placed on ice. The denatured probe was then added to the prehybridization solution
to a concentration of 10 ng/mL and replaced with hybridization solution overnight containing the
labeled probe. Sufficient hybridization solution is necessary to incubate at a ratio of at least 10
mL per 100 cm² filter area. The prehybridization solution was then discarded and replicated with
hybridization solution containing the labeled probe. The hybridization solution was recovered
and stored frozen. Once used again, the probe was denatured by heating at 95°C for 10 minutes
12
- 14. before use. The membrane was washed twice, 5 minutes per wash in Wash Solution #1
consisting of 2X SSC and 0.1% SDS at room temperature using disposable petri dishes. The
shaker incubator was used set at a slow speed to ensure no spilling occurred. The membrane was
then washed two more times for 15 minutes per wash with Wash Solution #2 consisting of 0.5X
SSC and 0.1% SDS. For probes longer than 100 bases, the wash needs to be performed at 60°C.
All following incubations were performed at room temperature. The washed membrane
was incubated in Wash Solution #3 consisting of 0.1 M Maleic Acid, 0.15 M NaCl pH 7.5, 0.3%
Tween20 for 1 minute. A freshly washed dish was used to block the membrane by agitating it in
Blocking solution (0.1 M Maleic acid, 0.15 NaCl, pH 7.5, 1% blocking reagent) for 30 minutes.
It was ensured that the filter was covered continuously during agitation. Towards the end of the
30 minutes, an antibody solution was prepared by diluting antiDIGalkaline phosphatase
1:5,000 in Blocking Solution by mixing 3 µL of antiDIG alkaline phosphatase in 15 mL
Blocking Solution for a final concentration of 150 mU/mL. This was mixed by gentle inversion.
The blocking solution was poured off after the 30 minutes and the membrane incubated with
gentle agitation for 30 minutes in the antibody solution created previously. The membrane was
transferred to a new dish and washed twice for 15 minutes each in a 20 mL Wash Solution #3 to
remove unbound antibodies. The membrane was equilibrated in 5 mL of detection buffer
consisting of 0.1 M TrisHCl, pH 9.5, 0.1 M NaCl for two minutes. The color substrate solution
was prepared by mixing 200 µL NBT/Xphosphate solution in 5 mL of detection buffer. The
color substrate solution was added to the substrate and in a plastic box in the dark. The
membrane was monitored for color development by being exposed to light for short periods.
Once spots were detected on the membrane results were documented by photograph.
Results
The products of the Cas9 PCR reaction and the restriction digest of the pcDNA
3.1/V5HisC plasmid were validated using agarose gel electrophoresis Figure 1a. The migration
distance of the restriction digest was compared to the DNA standard containing pieces of DNA
of known base pair lengths using the molecular marker in lane one and the restriction digest in
lane five of the agarose gel (Figure 1a). Using Excel, the distance the DNA standard fragments
13
- 18. The first step in confirming that the cells contained the pcDNA3.1/V5HCCas9 plasmid,
was isolating the plasmid using selective precipitation as described by the plasmid miniprep
protocol. Plasmids were isolated from three experimental cultures and one control culture, each
in a total volume of 50 µl. The purity and absolute amounts of plasmid pcDNA3.1/V5HCCas9
DNA recovered were found using ultraviolet spectroscopic analysis. Absorption readings were
taken at 260 nm (A260) and 280 nm (A260). The ratio of nucleic acids/protein, which determines
the purity of the sample, was calculated using A260/A280. The purity of the samples was
calculated for sample one and the control only, sample two and sample three had negative
absorption readings and could not be used. The absorption reading for sample one of the
pcDNA3.1/V5HCCas9 plasmid was A260 = 0.537 and A280 = 0.518. The ratio of nucleic acids to
protein, 0.537/0.518, was 1.04. The absorption reading for control was A260 = 0.034 and A280 =
0.008. The ratio of nucleic acid to protein, 0.034/0.008, was 4.25. The concentration of plasmid
DNA was calculated using the equation A260 x dilution factor x 50 µg/ml. The concentration was
used to calculate the absolute amount of DNA by multiplying the concentration by the total
volume recovered from the plasmid miniprep. The absolute amount of the
pcDNA3.1/V5HCCas9 plasmid from sample one was 26.85 µg and the absolute amount of the
control plasmid was 1.7 µg.
After determining the purity and amount of DNA in the samples, the plasmid was
linearized using the restriction digest Eco RI and run on an agarose electrophoresis gel along
with a marker, a DNA standard with a ladder of known base pair lengths. The gel (Figure 4a)
did not have a visible marker and the DNA that was visible on the gel was not distinguishable.
Using the ideal gel (Figure 4b) the length of the pcDNA3.1/V5HCCas9 plasmid digest was
calculated using regression analysis, (Figure 4c). The length of the pc3.1DNA/V5HCCas9
plasmid digest was 8256 base pairs. This was calculated by taking the antilog of the equation y=
0.0815x +5.2208, with x being the distance migrated (10.5mm) on the agarose gel.
Figure 4a (viewed on left) is the actual results
obtained in the laboratory. No molecular weight
marker was visible and DNA was indistinguishable
on the gel.
17
- 21. acid/protein ratio, which the A260/A280 ratio represents, is between 1.61.8 and neither the Cas9
product nor the plasmid restriction digest fall within this range. In addition, using concentrations
of the Cas9 PCR reaction and the restriction digest from the nanodrop spectrophotometer, the
absolute amounts of Cas9 DNA and pcDNA3.1/V5His C plasmid DNA were calculated. The
total amount of Cas9 DNA recovered was 20 ng compared to 5 ng of the pcDNA3.1/V5His C
plasmid DNA. The lack of purity may have resulted from improper extraction methods, which
would also explain the low amount of pcDNA3.1/V5His C plasmid DNA, during the
QIAQUICK DNA extraction. In addition, the unsuccessful PCR amplification accounts for the
low quantity of the Cas9 DNA.
The recombinant pcDNA3.1/V5HCCas9 plasmid was produced utilizing Gibson
cloning. The use of Gibson cloning allowed for the recombination of the Cas9 gene and the
pcDNA3.1/V5HisC using homologous recombination (4). This process reduces the need for
restriction digests, in addition to simplifying the process of making recombinant DNA. The
recombinant plasmid was then transformed into competent E. coli cells. These cells were plated
on LB plates that included ampicillin in the media. E. coli is not naturally resistant to ampicillin
but the recombinant plasmid contains the ampr
gene that confers ampicillin resistance, and if
transformation was successful the cells would grow colonies on the ampicillin plates when
incubated overnight. To ensure the efficacy of the ampicillin plates a negative control,
containing no plasmid DNA was also incubated overnight. The efficacy of the competent cells
was tested by plating competent cells transformed with pcDNA3.1/V5His C plasmid DNA onto
an ampicillin plate and incubated overnight. The negative control showed no growth, confirming
that the ampicillin plates were effective. However, although the positive control plates did show
growth there were very few colonies. The experimental plates showed no growth. The
transformational efficacy of the plate containing 50 µl of the positive control was 200
transformants/µg. The plate containing 200 µl of the positive control had a transformational
efficacy of 320 transformants/µg. The transformation efficiencies are well below typical
transformation efficiencies, with even general transformation efficiencies being 106
transformants/µg. The low numbers may have resulted from lack of incubation, and extended
incubation times may be necessary in the future.
20
- 22. The next step taken was to run an agarose gel with the recombinant
pcDNA3.1/V5HCCas9 plasmid. The recombinant plasmid was linearized using the Eco RI
restriction enzyme. Three samples of the purified recombinant plasmid, a control, and a standard
DNA ladder were loaded into separate wells on the agarose gel (Figure 4a). After running the
gel, there was no visible standard DNA ladder, and the product did not produce clear bands for
analysis. The ladder did not appear on the gel because cyber green was not added to the loading
buffer containing the standard. The streaking that occurred on the gel could be a result of DNA
degradation of the samples. In addition, the E. Coli cells transformed with the recombinant
plasmid, used to recover the recombinant pcDNA3.1/V5HCCas9 plasmid, were not grown
overnight in culture and were grown for a shorter amount of time than the protocol required.
This could be the reason for the poor recovery in the plasmid DNA, since replication of the E.
coli cells is needed to produce copies of the pcDNA3.1/V5HCCas9 plasmid. Using the ideal gel
(Figure 4b) a standard curve was generated (Figure 4c) to calculate the regression line y =
0.0815x + 5.2208. The length that the recombinant plasmid DNA traveled on the gel was
entered as the xvalue in order to solve for y, the antilog of the y value corresponds to length of
the sample containing the recombinant pcDNA3.1/V5HCCas9 plasmid. The length of
pcDNA3.1/V5HCCas9 plasmid was calculated at 8256 base pairs and the actual length of the
plasmid is 9634 base pairs. The difference in lengths is likely from measurement errors in either
creating the regression curve or measuring the distance travelled by the recombinant plasmid.
The last step to confirm that the recombinant pcDNA3.1/V5HCCas9 plasmid was
successfully cloned was performing the dot blot. The dot blot is a powerful tool for confirming
the presence of a specific sequence of DNA. After denaturing the plasmid to single strands of
DNA, the denatured recombinant plasmid was spotted on and UVcrosslinked to a nylon
membrane. In the previous step, there was a failure in recovering the pcDNA3.1/V5HCCas9
plasmid, and samples from another laboratory section were used to perform the dot plots. The
membrane was hybridized using the DIG probe. The DIG probe is complementary to the Cas9
gene and the denatured plasmid is able to attach to the complementary strand. The probe is
labeled with dioxigenin that allows for color detection of the probe. The two membranes were
treated with four samples of plasmid DNA, two positive controls, and a negative control. The
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- 23. positive control was used to ensure that the probe was successful in attaching to the target
sequence. The negative control is also used to ensure that the probe attached to the
complementary sequence. If there is no color detection on the negative control, but there is color
detection on the positive control, specific binding occurred. It can then be concluded that color
detection on the experimental dot blot is the result of specific binding, and the Cas9 gene is in
that experimental plasmid. Conversely, if the negative control detects color, no conclusions can
be drawn, because color detection on the negative control is a result of nonspecific binding. Two
dot blots were performed and one was successful (Figure 5a), while the other was unsuccessful
(Figure 5b). On the successful dot plot, color detection was seen in all four experimental
samples, and in these samples cloning of the pcDNA3.1/V5HCCas9 plasmid was achieved. On
the other membrane (Figure 5b) there was color detection on all of the experimental samples
and the control, so no conclusions can be made about these samples. The color detection on the
negative control could be a result of nonspecific binding due to an improper hybridization
temperature or the stringency of the wash being too low.
This experiment aimed to subclone the pcDNA3.1/V5HCCas9 plasmid using E. coli as
an expression vector. Amplification of the Cas9 plasmid was unsuccessful, recovery of the PCR
product resulted in only 20 ng of the Cas9 DNA. Additionally, only 5 ng of DNA was recovered
after DNA purification of the pcDNA3.1/V5HisC plasmid. Gibson cloning was run using low
concentrations of starting materials, and the E. coli transformed with the recombinant
pcDNA3.1/V5HCCas9 plasmid from the Gibson assembly were not grown for the proper
amount of time that would allow for saturated cultures. Without high quantities of recombinant
plasmid from the Gibson cloning, and saturated transformed E. coli vectors, there was not quality
recovery of the recombinant pcDNA3.1/V5HCCas9 plasmid. The results of the dot plot, with
plasmids recovered from another lab section, did show that at least four of the experimental
transformed E. coli vectors produced the recombinant plasmid and the plasmid contained the
Cas9 gene. This result is consistent with the main objective of this lab series, subcloning the
pcDNA3.1/V5HCCas9 plasmid into a new expression vector.
Future applications are an important reason for performing such experiments. Future
applications include, but are not limited to, engineering and programming RNAguided DNA
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