Team:St Andrews/Experiments

Experiments

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• Miraprep & miniprep
• Plasmid/gBlock double digest
• Gel purification
• Ligation
• Transformation
• Cell culture

Miraprep

Background

The Miraprep is a method that uses commercial Miniprep kits for the purification of DNA using a silica column, to get results comparable to industrial purifications. The process essentially runs 5 Minipreps at once resulting in a greater yield of stable plasmid. This step was used to purify large quantities of unaltered vector DNA for digestion, gel purification, and ligation with our DNA g-blocks. The protocol was adapted from Pronobis et al. 2016.

Aim

To purify recombinant plasmid DNA from transformed E. coli cells in high concentrations for sequencing and further transformations.

Materials

• QIAprep Spin Miniprep Kit (1)
• 96% ethanol
• 50mL Falcon tube
• 50mg/mL RNase
• Centrifuge
• 1.5mL microcentrifuge tubes
• Microcentrifuge
• 15mL Falcon tube
• Distilled Water

Protocol

The QIAprep spin miniprep kit was collected and it was ensured that the DNA wash buffer was mixed with ethanol. Following this, a 50mL overnight culture was transferred to a 50mL Falcon tube and spun down at 3230xg for 20 minutes* at 4°C. The supernatant was discarded, and the pellet was resuspended in 2mL of resuspension buffer (P1) with 50ug/mL RNase – fresh RNase was required each time. The resuspended 2mL was then aliquoted into 4 seperate 1.5mL centrifuge tube, each with 0.5mL of the resuspension*. Lysis buffer, 0.5mL, was added to each aliquot which was then inverted 4 times, and then left to incubate for 3 minutes at room temperature.Neutralisation buffer, 0.5mL, was added to each aliquot and the microcentrifuge tube was inverted 4 times before the tubes were then spun at 13,000xg for 10 minutes at room temperature. The supernatant was collected in a 15mL tube and the pellets were discarded. 1x volume (6mL) of 96% ethanol was added and the solution was mixed thoroughly for 5 seconds.

The solution was then loaded onto 5 spin columns in 3 sequential 700uL aliquots and spun at 13,000xg for 30 seconds before flow through was discarded. The columns were washed twice with 500uL of wash buffer, being spun after each wash at 13,000xg for 30 seconds and the flow through discarded. The empty columns were spun once more at 13,000xg for 1.5 minutes to remove any remaining buffer. Following this, the old collection tube was discarded, and each column placed into a new spin tube. The DNA was eluted from the column by adding distilled water, 35uL. This was left to incubate for 2 minutes at room temperature, and spun at 13,000xg for 2 minutes. The eluted DNA was collected in one tube and measured on the Nanodrop before the microcentrifuge was clearly labelled and stored at -20°C. * denotes an adaptation to the published procedure.

Miniprep

Background

The purpose of the Miniprep is to purify plasmid DNA from a cell culture for further use. Minipreps typically produce ~50µL of purified DNA. This technique was used when lower levels of plasmi were required to avoid using a Miraprep.

Materials

• QIAprep Spin Miniprep Kit (1)
• 1.5mL microcentrifuge tubes
• Microcentrifuge
• Distilled Water
• Centrifuge
• 15mL Falcon tube

Protocol

~5mL of cells taken from an overnight culture were spun down in a 15mL Falcon tube in a centrifuge at 4,200xg for 10 minutes and the supernatant discarded. The pellet was then resuspended in 250µL of buffer P1 (ensuring RNase was added beforehand) and transferred to a 1.5mL microcentrifuge tube. Following this, 250µL of buffer P2 was added and the tube was inverted 6 times to ensure thorough mixing. Buffer N3, 350µL, was then added and the mixing step above repeated immediately. The tube was then spun down in a microcentrifuge at full speed for 10 minutes. The supernatant from this process was then added to a QIAprep spin column and spun down for 60 seconds. The column was then washed with 0.5mL of buffer PB and centrifuging again for 60 seconds. The column was further washed by adding 0.75mL and centrifuged for a further 60 seconds. The flow through was discarded and centrifuged once more for 60 seconds before the flow through was discarded. Finally, 50µL of distilled water was added to the centre of the column and left to incubate for 1 minute. The column was then spun down at full speed for 1 minute.

Digest of gBlocks

Background

The DNA sequences for the proteins we desired to express were constructed by designing plasmids containing everything required for cloning, digesting, expressing, and purifying. These protein sequences were optimised for E. coli and ordered via Integrated DNA Technologies. In order for these gBlocks to be inserted into our chosen plasmids (varying between pEHISTEV, pUC19, ad pSB1C3), the ends had to be cleaved to allow ligation into the vector. Over the course of the project varying enzymes were used depending on the vector (table 1). The following protocol was written up for an NdeI and PciI double digest.

Table 1. Restriction enzymes used in each vector

Materials

• Water bath at 37°C
• 1.5mL microcentrifuge tube
• NEB NdeI
• NEB PciI
• Distilled water
• Buffer 3.1
• 10ng/µL of gBlock

Protocol

In a 1.5mL microcentrifuge tube, a volume of 25µL of gBlock was mixed with 3µL of 3.1 buffer and 2µL each of NdeI and PciI. This was then kept in a water bath for 2 hours at 37°C. Following completion, the digest could then be used in a ligation with purified and digested plasmid.

Digest of Plasmids

Background

The plasmids used in our cloning were pSB1C3 for submission of parts, pEHISTEV which contains a promoter and so was used for overexpression, and pUC19 which was chosen for ease of cloning in preliminary rounds. The restriction sites used in each plasmid is shown in table 1. The following protocol was written up for an NdeI and PciI double digest of pUC19.

Materials

• Water bath at 37°C
• 1.5mL microcentrifuge tube
• NEB NdeI
• NEB PciI
• Distilled water
• Buffer 3.1
• 2.5µg of gBlock

Protocol

In a 1.5mL microcentrifuge tube, 2.5µg of plasmid DNA was mixed with 3µL of 3.1 buffer, 2µL each of NdeI and PciI, and made up to 30µL distilled water with the enzyme being added last. This was then kept in a water bath at 37°C for 2 hours. Following completion, the digest could then be purified via gel electrophoresis and used in a ligation with gBlock constructs cleaved with the same enzymes. A single digest was also run for each of the restriction enzymes.

Note

It is noted in the original paper on the Miraprep (Pronobis, Deuitch and Peifer, 2016) that over time the RNase in buffer P1 can degrade. This note was not originally caught by our lab team and as a result, Mirapreps of pSB1C3 and pEHISTEV likely gave us samples with higher levels of RNA and therefore deceptive DNA concentrations when analysed via the nanodrop spectrometer at OD260. This was identified on gels run after the digest which showed little to no DNA despite 2.5µg apparently used. As a result, the plasmid volume used was increased 10-fold. Due to our inability to appropriately identify the concentration of DNA in our plasmid stock, the exact mass of DNA used is unknown. Following purification, yields of roughly 20ng/µL were given.

1% Agarose Gel and Gel Purification

Background

The 1% agarose gel is used to separate DNA fragments based on size. In this example it was used to separate the cut plasmid to be used for insertion from the smaller fragment that has been removed. The single digests were also both run on the gel as a control. A QIAquick Gel Extraction Kit was then used on the removed band to purify the DNA.

Materials

• Gel tray and lane fork
• Agarose powder
• 1x TAE buffer
• SYBR Safe
• Powerpac
• Microwave
• QIAquick Gel Extraction Kit
• DNA loading dye

Protocol

A 1% agarose gel was prepared by mixing 0.5g of agarose with 50mL of 1xTAE buffer. This was heated in a microwave in 30 second increments to ensure the solution did not boil and was handled with care at all times. This was left to cool until it was possible to touch it and 5µL of SYBR Safe dye was added. Following this the gel was poured into the gel tray containing the gel fork and was left to set – ensuring no air bubbles were present. Upon setting of the gel and removal of the lane fork, 6µL of 6x molecular weight ladder was added to the first lane and the double and single digests were each mixed with µL of loading dye and added into separate columns. This was then run for half an hour at 90V. Following the running of the gel, it was visualised under an LED light box – thanks to the use of SYBR Safe, no UV box was necessary.

Using a clean scalpel and under the LED light, the desired band of DNA for our cleaved plasmid was identified and cleaved from the gel – cutting as close to the band as possible and removing excess gel. The band was put into a 1.5mL microcentrifuge tube and weighed to record the mass of the band. Using 100mg = 100µL, 3 volumes of buffer QG was added to the microcentrifuge tube and the tube was incubated at 50°C until the gel had fully dissolved – vortexing every 2-3 minutes to aid the process. Following this, 1 volume of isopropanol was added and the sample was transferred to a QIAquick spin column within a 2mL collection tube. The column was filled and spun down in a microcentrifuge at 10,000xg or 13,000rpm for 1 minute, the flow through discarded, and then loaded again until all of the sample was loaded on the column. Buffer QG, 0.5mL, was added to the column and centrifuged as above with the flow through discarded. Buffer PE, 0.75mL, was added to the column and the column was left to stand for 5 minutes before being centrifuged as above with the flow through discarded and the column was centrifuged for an additional minute. The column was then placed in a clean microcentrifuge tube and 30µL of water was added. The column was left for 1 minute before being spun as above for a final time. The final sample was then labelled and analysed via nanodrop.

Ligation of gBlocks and Plasmids

Background

The process of ligation is necessary to join the digested plasmids and gBlocks together to allow transformation of bacterial cells. By using T4 DNA ligase we were able to ligate our desired gBlocks with their vectors in as little as 5 minutes.

Materials

• 30-40ng of plasmid
• DNA construct
• Distilled water
• 2.5µg of gBlock
• Ice bucket
• 1.5mL microcentrifuge tube
• NEB 10x T4 ligase buffer
• NEB T4 DNA liagse
• Water bath at 42°C

Protocol

A 1.5mL microcentrifuge was filled with the plasmid, 6 µL of construct, 1 µL of 10x T4 ligase buffer, and finally 1 µL of T4 DNA ligase. A negative control was run with the same set up but with 6µL of water in place of construct. Once the microcentrifuge tubes were filled, they were left at room temperature for 5 minutes. Following this a transformation would be carried out.

Bacterial Cell Transformation

Background

This protocol is a standard heat shock method for the insertion of recombinant plasmids into target cells. Heat shock takes advantage of a steep temperature gradient to increase the permeability of cell membrane, resulting in the uptake of the new DNA.

Materials

• Competent E. coli DH5α cells
• Sterile spreaders
• Distilled water
• Sterile LB
• Ice bucket
• Stationery 37°C incubator
• 1.5mL microcentrifuge tube
• Recombinant plasmid
• Orbital shaking incubator
• Water bath at 42°C
• Sterile ampicillin plates
• Microcentrifuge

Protocol

Competent cells were transferred from the -80°C into an ice bucket and left to defrost until liquid (20-30 minutes). In this time, 1µL of plasmid was pipetted into a 1.5mL microcentrifuge tube and was added to with 50µL of the competent cells. The microcentrifuge tubes were then transferred onto ice for 20 minutes. Following the 20 minute period, the tubes were placed in a water bath at 42°C for 45 seconds before being immediately returned to ice. After 5 minutes on the ice, 900µL of sterile LB was added to each tube and they were taped side down in an orbital incubator at 37°C for 1 hour.

After incubation, the tubes were spun down in a microcentrifuge for 1 minute at full speed, 800µL of supernatant was discarded and the cells were resuspended in the ~100µL remaining. The resuspended cells were then pipetted onto a sterile agar ampicillin plate, spread evenly using sterile spreaders, and stored agar side up in a 37°C stationery incubator overnight.

Cell Culturing

Background

Overnight cell cultures are used to produce many bacteria for further isolation of modified DNA.

Materials

• Inoculated and incubated agar plate with specific bacteria
• Sterile LB
• Sterile culture flask
• Liquid antibiotic
• Orbital shaking incubator at 37°C

Protocol

A sterile culture flask was filled with 10mL of sterile LB was mixed with 10µL of the specific antibiotic from a pre-prepared stock. Into this, a pipette tip was ejected after touching it against an isolated round colony. In some cases, the ideal site could be identified by colour depending on the proteins expressed. This was left in an orbital shaking incubator at 37°C overnight.

Incubation for Expression of Construct 9

Background

To observe the expression of mNeongreen in our bacterial cells, we had to culture them to specific optical densities in order to observe the differences in absorbance following the addition of IPTG which activates expression.

Materials

• Bacterial stock solution
• Sterile LB
• Sterile culture flasks
• Chloramphenicol
• Orbital shaking incubator at 37°C
• Spectrophotometer

Protocol

Absorbance of stock solution was measured at 600nm at the following concentrations:

• 4x 250µL stock, 750µL LB, A=1.37
• 10x 100µL stock, 900µL, A=.39
• 20x 50µL stock, 950µL LB, A=.26
• 40x 25µL stock, 975µL LB, A=.15

Two flasks were prepared at 40x dilution**:

• 48.75ml LB
• 1.25ml stock
• 50µL chloramphenicol (25 mM)

Flasks are left to incubate until absorbance reached 0.6

After, 50µL of IPTG was added to a single flask with the other left as a negative control. The culture was then left overnight.

** dilution does not matter as much as absorbance, dilutions should be made from the stock until A is approximately 0.1

Western Blot

Background

A Western blot relies initially upon the use of an SDS-PAGE to separate proteins by mass allowing identification of proteins. Following this a blot takes place to transfer the proteins to a membrane with high protein affinity. On this membrane, antibodies specific to the protein of interest can be incubated. A secondary antibody that binds to the original can then be added that has a signal response – often fluorescence. This reporting method allows a rough quantification of the proteins present as well as a more concrete identification of the specific protein. This was used in our investigation to observe whether our protein was being expressed and where in the cell the protein was being expressed.

Materials

• Pellet samples of each construct
• SDS sample buffer
• Heating block at 95°C
• 8% acrylamide gel
• Blotting unit
• Prestained protein ladder
• Milk powder
• Secondary antibody
• Sponges, 3MM paper, protein membrane
• PBS
• Ponceau red
• V5 tag
• Construct 9 culture
• Spectrophotometer
• Powerpac
• Gel knife
• Distilled water
• Primary antibody
• Roller
• Tween 20
• 50mL Falcon tube
• Gel imager

Protocol

The pellets of each construct in microcentrifuge tubes were resuspended in 100µL of reducing agent (5x SDS and water) and heated at 95°C for 10 minutes. A negative control was produced from just construct 9 expressing cells as construct 9 doesn’t have a V5 tag. A positive control was produced containing a V5 tag only. Following the heating of all samples, a premade acrylamide gel was removed from its wrapping, rinsed with deionised water, and inserted into a minicell. The minicell was then filled from the inner chamber out with running buffer. Into the wells of the gel, 5µL of the ladder, 20µL of the positive control, and 25µL of the negative control and all other solutions were loaded. The SDS-PAGE was then run at 200V for 45 minutes and then gradually continued until the dye front reached close to the end of the gel.

While the gel was running, the sponges, 3MM paper, and the protein binding membrane were soaked in running buffer – with the membrane soaking independently flanked by tissue paper. The gel was removed from its case and trimmed down before being rinsed with distilled water. The transfer stack was then constructed with the sponges on the outer layer, the 3MM paper inside the sponges, and the membrane and the gel in between the paper. This was then inserted into the blotting unit and ran for 1.5 hours at 100V with an ice pack in the running before to prevent melting.

Following this, the membrane was removed from the casing and inserted into a 50mL Falcon tube with 5% milk solution as a blocking buffer and left on a roller for 2 hours. The membrane was then transferred into a second Falcon tube containing 5% milk solution and 5mL primary antibody solution. This was incubated for 1 hour on rotating rollers. Next, the membrane was rinsed twice with 50mL PBS-T and transferred to another 50mL falcon tube containing 40mL PBS-T for a further 5 min incubation with shaking (room temperature). The membrane was rinsed twice again with 40mL PBS-T with the 5 min incubation step in between.

After the PBS-T was removed, 5mL of the secondary antibody solution was added to the falcon tube containing the membrane and incubated in the dark for an hour on rotating rollers. Then, the washes described after the primary antibody solution were then repeated in the exact manner. 50mL PBS was then used to rinse the membrane and the signal was visualised using an Odyssey scanner.