Genomic Editing by Subcloning of the Cas9 gene from

pCAMBIA­Cas9 snRNA Plasmid into pcDNA3.1/V5­His C

Expression Vector

Margaret Sullivan, Alexandria Rector, Mari Moe Dagslet

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

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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

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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

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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

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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,

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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

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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

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adjacent sequence and seals the backbone. This entire process produces a double­stranded fully

sealed DNA molecule.

The first step of the Gibson Cloning was to calculate the number of pmols of each

fragment for optimal assembly. A total of 0.03­0.2 pmols of DNA is recommended when 1­2

fragments are assembled into a vector. Efficiency decreases as the number or length of fragments

increase. Amount of pmols is easily calculated by using the formula pmols = (weight in ng) x

1000 / (base pairs x 650 daltons), 50 ng of 5000 bp dsDNA is around 0.015 pmols while 50 ng of

500 bp dsDNA is around 0.15 pmols. The mass of each fragment is measured using the

Nanodrop or agarose gel electrophoresis.

An assembly protocol was created using amounts as stated in the table below:

Assembly* Assembly** Control

Recommended DNA Ratio

vector:insert = 1:2 vector:insert = 1:1

Total No. of Fragments 0.03­0.2 pmols X μl 0.2­0.5 pmols 10 μl

2X Assembly Mix 10 μl 10 μl 10 μl

Deionized H2O 10­X μl 10­X μl 0

Total Volume 20 μl 20 μl 20 μl

Table taken out of Bio 351 lab manual, Dr. Read 2015.

This assembly was incubated at 50°C for 60 minutes, utilizing one­step isothermal DNA

assembly. The process first described by Gibson, D.G. et. al, allows for recombination of the

cassettes into the vector at a constant temperature, rather than the cycling previously used,

making the process simpler without reducing efficiency (3). After incubation, the samples were

stored on ice for subsequent transformation.

Bacterial Transformation

Bacterial transformation was completed following Gibson assembly to introduce a

foreign plasmid into bacteria and use that bacteria to amplify the plasmid. The bacterial

transformation was completed by using 3 different aliquots of competent cells to set up 3

transformation reactions. One with a positive control and 100 ng of pCDNA3.1/V5­His c, one

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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

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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.

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Plasmid DNA Digestion

Digestion of plasmid DNA was the next process to be completed. This process consists of

cutting the recombinant plasmid pcDNA3.1/V5HC­Cas9 with the restriction enzymes, Eco RI

and KpnI. A double digest with these restriction enzymes will release the Cas9 gene from the

recombinant plasmid. The digested DNA will be run on an agarose gel and will migrate as

distinct bands on the gel due to being cut into two different­sized fragments.

It was incredibly important during this part of the experiment to be extra careful with the

restriction enzymes. Restriction enzymes are stored in concentrated glycerol for stability, these

enzymes can be easily denatured in adverse conditions and are expensive to replace. For these

reasons, they must be kept on ice and returned to the freezer rapidly. Gloves must always be

worn when handling these enzymes to prevent contamination.

A reaction mix was assembled on ice with 2 µL of 10X reaction buffer, 17.5 µL of

plasmid DNA solution (0.75 µg), and 0.5 µL of Eco RI for a total of 20.0 µL. A total of four

microfuge tubes containing the reaction buffer, plasmid DNA, and restriction enzyme were

assembled. It was necessary to add the components to the microfuge tube in the order listed to

ensure the reaction did not start earlier than intended. The restriction enzymes cleave plasmids at

specific sites, and the restriction enzyme was used in this lab in order to linearize the DNA. The

10X reaction buffer provided the salt conditions necessary for the restriction enzyme to function.

The reactions were centrifuged 5­10 seconds to combine the components and begin the reaction.

The tubes were placed in a 37°C water bath and incubated for at least one hour to complete the

reaction.

Dot Blot

The Dot Blot technique was the final process in the steps of this experiment. The Dot

Blot Technique is the process of nucleic acid hybridization used to identify specific DNA and

RNA molecules and quantify the relative amounts of nucleic acids with known homology.

Throughout this process, it is necessary to wear gloves to protect your hands and prevent

contamination. It is also preferable to use forceps when manipulation nylon membrane to prevent

contamination. This process began by cutting a strip of nylon membrane to the desired size and

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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

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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

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migrated was plotted versus the log of the length in base pairs and a standard curve was

calculated (Figure 1b). The molecular weight restriction digest was calculated using regression

analysis. The value calculated was 6471 base pair in length. This was calculated by inserting

distance the restriction digest traveled from the well (17mm) into x of the regression line

generated on the standard curve, y= ­0.0501x+4.6627, and taking the antilog of this value.

Figure 1a on the left is the agarose gel containing the standard DNA ladder on the far left and the restriction digest in the lane denoted by the yellow arrow. Figure 1b on the right is the graph of the standard curve generated and the regression line used to calculate the size of the pcDNA3.1/V5­His C plasmid restriction digest by plotting the distance migrated (mm) versus the log of the marker length in base pairs.

No PCR product was visible on the agarose gel, and an ideal gel (Figure 2a) was used to

calculate the length of the PCR product in the same manner described for the restriction digest.

The standard curve generated in Excel for the PCR product (Figure 2b) was used to calculate the

length of the Cas9 plasmid PCR product. The length of the Cas9 PCR product was calculated to

be 5446 base pairs is length. The value was calculated using distance migrated by the Cas9 PCR

product (17mm) on the standard gel, and plugging this value in for x in the regression equation

y= ­0.0475x +4.5436.

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Figure 2a Ideal agarose gel with Cas9 PCR product used to generate the standard curve in Figure 2b Figure 2b on the right is the graph of the standard curve generated and the regression line used to calculate the size of the Cas9 PCR product by plotting the distance migrated (mm) versus the log of the marker length in base pairs.

The Cas9 PCR product and 3.1/V5­HisC plasmid were purified using QIAquick DNA

purification. A nanodrop spectrophotometer was used to find the purity and concentration of the

PCR product and plasmid digest. The Nanodrop readings are reported in absorption at 260 nm

over absorption at 280 nm (A260/A280). These values correspond to the wavelengths that nucleic

acids and proteins absorb, and the A260/A280 reading represents the ratio of nucleic acids to

protein. The A260/A280 reading for the Cas9 PCR product was 3.23 and the concentration was 0.4

ng/µl. Using the concentration and the total volume of Cas9 PCR product, 0.4 ng/µl x 50 µl = 20

ng, the absolute amount of Cas9 PCR product DNA obtained after purification was 20 ng. The

A260/A280 reading for the 3.1/V5­HisC plasmid was ­0.18 with a concentration of 0.1ng/µl. The

absolute amount of 3.1/V5­HisC plasmid DNA obtained after purification was 5 ng, 0.1 ng/µl x

50 µl = 5 ng.

Gibson cloning was used to make the recombinant pcDNA3.1/V5HC­Cas9 plasmid from

the purified Cas9 and 3.1/V5­HisC plasmid DNA. The assembly mix contained 5 µl of Cas9 and

5 µl of 3.1/V5­HisC plasmid. The ratio of Cas9 DNA to 3.1/V5­HisC plasmid DNA was 4:1,

found by multiplying the volumes added to the assembly mix by the concentrations reported by

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the nanodrop. In order to clone the recombinant plasmid, it was introduced into E. coli cells

using bacterial transformation. Bacterial transformation was accomplished by using competent

cells previously prepared for this lab, and combining the cells with the recombinant plasmid

followed by quick heat shock. The recombinant plasmid contained the ampr gene, the gene

responsible for ampicillin resistance. In order to confirm the E. coli cells were successfully

transformed with the recombinant plasmid the cells were plated on ampicillin containing media.

The efficacy of the ampicillin was tested using the negative control while the competence of the

cells used to transform the E. coli was confirmed using a positive control. The negative control

did not grow any cells, confirming that the ampicillin was effective, and the positive control did

have growth confirming that the cells were competent (Figure 3).

Figure 3. Actual results of LB plate

containing ampicillin with positive control

competent cells. 50 µl of positive control

viewed on the left contains one colony. 200

µl of positive control on the right contains 8

colonies. Colonies are the visible yellow

spots.

One colony was counted for the plate containing 50 µl of the positive control, while eight

colonies were counted on the plate containing 200 µl of positive control. There was no growth

on the experimental plates containing the E. coli cells transformed with the recombinant

pcDNA3.1/V5HC­Cas9 plasmid. The efficiency of transformation (number of colonies/µg of

added DNA) was calculated for the positive control competent cells using the equation:

number of colonies/ng of DNA x dilution factor x 1000 ng/µg.

The 50 µl positive control plate (1 colony/100ng of DNA x 20 DF x 1000 ng/µg) had a

transformation efficiency of 200 transformants/µg and the 200 µl positive control plate (8

colonies/100ng of DNA x 5 DF x 1000 ng/µg) had a transformation efficiency of 320

transformants/µg.

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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.

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Figure 4b (viewed on right) is the ideal gel with the molecular weight marker in well 1 on the far left. Length of the recombinant pcDNA3.1/V5HC­Cas9 plasmid digest was calculated using regression analysis.

Figure 4c graph of the standard curve generated and the regression line used to calculate the size of the recombinant pcDNA3.1/V5HC­Cas9 plasmid by plotting the distance migrated (mm) versus the log of the marker length in base pairs.

Performing a dot blot was the final step in confirming successful cloning of

pcDNA3.1/V5HC­Cas9 plasmid. Two membranes were prepared, each dotted with four

experimental pcDNA3.1/V5HC­Cas9 plasmids, two positive controls, and a negative control.

After hybridization with probes containing sequences complementary to the

pcDNA3.1/V5HC­Cas9 plasmid, the membranes were developed in the dark until color started to

develop. All of the experimental dots and the positive controls developed as shown in Figure

5a, but the negative control did not, and therefore the experimental samples contain the

pcDNA3.1/V5HC­Cas9 plasmid and cloning was successful. In Figure 5b all of the dots

developed, including the negative control, and because of this no conclusion can be made about

the plasmid samples on this membrane. Figure 5a. Dot blot containing experimental samples with pcDNA3.1/V5HC­Cas9 plasmid in the top four boxes all showing colored dots. The bottom row contains visible dots in the positive controls (two bottom right boxes) and the negative control (to the left of the positive controls) showed no color..

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Figure 5b. Dot blot containing samples from another lab section. The top four boxes starting from the left hand side contain the experimental recombinant plasmid. Boxes on the bottom right with positive (far right two boxes) and negative control (left box) appeared showing an inconclusive result.

Discussion

The first step in cloning the Cas9 gene into the pcDNA3.1/V5­His C plasmid was to

amplify the Cas9 gene using a PCR reaction. The product of the PCR reaction and a restriction

digest of the pcDNA3.1/V5­His C plasmid were then run through an agarose gel using

electrophoresis. A DNA standard ladder of known base pair lengths was also run in order to

create a standard curve, which was then used to calculate the lengths of the Cas9 PCR product

and the restriction digest. The restriction digest was successful, but the PCR product was not

visible on the gel and the ideal gel was used to calculate the length of the PCR product (Figure

1a & 1b). From the antilog of standard curve for the restriction digest, y= ­0.0501x+4.6627, the

length of the pc3.1DNA/V5­HisC restriction digest was calculated to be 6471 base pairs. The

actual length of the plasmid is 5498 base pairs. The difference between the length calculated and

actual length is likely a result of measuring errors, as the measurements were taken using a

photograph and not the actual gel. The same process was used to calculate the Cas9 PCR

product using the equation y= ­0.0475x + 4.6627, and taking the antilog of the y value. This

equation was generated using the ideal gels provided (Figure 2b). The calculated length of the

PCR product using this equation was 5446 base pairs, while the actual length of the Cas9 gene is

4136 base pairs. The difference in values is again likely due to errors in measurement. The

amplification of the PCR reaction was unsuccessful and could be a result of a high annealing

temperature. The number of cycles the PCR was run may need to be increased because of the

large size of the Cas9 gene being amplified.

The nanodrop spectrophotometer was used to estimate the purity of the Cas9 PCR

product and the pcDNA3.1/V5­His C plasmid restriction digest using the A260/A280 reading. This

reading is used to estimate the purity because the nucleic acids absorb wavelengths at 260 nm

and proteins absorb wavelengths at 280 nm. The Cas9 PCR product A260/A280 ratio was 3.23 and

the pcDNA3.1/V5­His C plasmid restriction digest A260/A280 ratio was ­0.18. An ideal nucleic

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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.

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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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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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endonucleases (1), using CRISPR to edit several different sites within the mammalian genome

(2), and eventually use CRISPR/Cas9 to conduct gene editing in human zygotes (5). Gasiunas et

al. found that altering the RNA within the Cas9­crRNA complex allows endonucleases to be

designed for in vivo and in vitro use. These revelations allow for the development of exclusive

tools for RNA­directed DNA surgery. Another use for the CRISPR­Cas9 method is encoding

multiple guide sequences into a CRISPR array for several site editing within the genome of

mammals (2). This again demonstrates varied applications and programmability of technology

using RNA­guided nuclease. Liang et al. began using this gene editing in human tripronuclear

zygotes, a step toward using this in viable human embryos. While this research exhibited

uncertain results, it is research that will be edited and utilized in the future. The CRISPR/Cas9

system can be used in various ways to edit, knock out, or replace specific genomic sequences.

References

Gasiunas et al. 2009. Cas9­crRNA ribonucleoprotein complex mediates specific DNA cleavage

for adaptive immunity in bacteria. PNAS, (Don’t know exactly how to cite this)

Cong et al. 2013. Multiplex genome engineering using CRISPR/Cas systems. Science, 339:

819­823.

Liang et al. 2015. CRISPR/Cas9­mediated gene editing in human tripronuclear zygotes.

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