Protocols

Plasmid Isolation

• Cell Harvesting: Bacterial cells containing the desired plasmid are grown in culture and then harvested. The cells are typically treated to disrupt their cell walls, releasing the plasmids into the solution.
• Cell Lysis and DNA Release: The harvested cells are subjected to lysis, breaking open their membranes and releasing cellular components. Plasmid DNA, along with other cellular components, is liberated into the lysate.
• Removing Cellular Debris: The lysate is centrifuged to separate the cellular debris and larger particles from the liquid containing the plasmid DNA.
• Selective Precipitation or Column Purification: Plasmid DNA can be selectively precipitated using alcohol, where the DNA molecules aggregate while other impurities remain in solution. Alternatively, column purification methods employ specialized matrices that selectively bind DNA while allowing contaminants to be washed away.
• Elution and Concentration: The purified plasmid DNA is eluted from the purification matrix or recovered from the precipitation process. It is then concentrated through ethanol precipitation or centrifugation.
• Quality Assessment: The extracted plasmid DNA is assessed for quality and quantity using techniques like spectrophotometry or agarose gel electrophoresis.

Kitfree Alkaline Lysate method:

Solutions 1, 2, 3 prepared for Alkaline Lysis Plasmid Extraction: 

1. Solution 1:
   1. 25mM Tris HCL (pH=8)
   2. 50mM Glucose 
   3. 10mM EDTA  
2. Solution 2:
   1. 0.2N (0.2M) NaOH
   2. 1% SDS 
3. Solution 3:
   1. 120mL 5M Potassium Acetate 
   2. 23mL Glacial Acetic Acid 
   3. 57 mL dH₂O  

TE: Tris EDTA buffer:

1. 10mM Tris
2. 1mM EDTA
3. Bring to pH=8.0

Protocol: 

• Grow 2 mL overnight cultures from single colonies of bacteria containing your plasmid of interest.
• Add 1.5 mL of the stock culture to a 1.75 mL microfuge tube.  
• Centrifuge in microfuge tube at 10,000 g for 30 sec. 
• Pour off the supernatant, being careful not to disturb the bacterial pellet. 
• Resuspend the pellet in 100 mL of cold Solution I. 
• Vortex the solution for 2 min or until all bacteria are fully resuspended.
• Add 200 mL of Solution II and invert the tube carefully 5 times to mix the contents. The contents will become clear and thicker as the proteins and DNA are denatured. Do not vortex at this stage or the genomic DNA will become sheared and will therefore contaminate purified plasmid DNA. 
• Incubate solution on ice for 5 min. 
• Add 150 mL of cold Solution III to each tube. 
• Mix by inverting several times. A white precipitate will be formed which contains bacterial proteins and genomic DNA. 
• Incubate tube on ice for 5 min. 
• Centrifuge the tube for 5 min at 12,000 g. 
• Collect the supernatant into a new tube by pipetting or carefully pouring. 
• Add 5 mL of 2 mg/mL RNase A to the supernatant in the new tube and incubate at 37°C for 5 minutes.
• Perform phenol-chloroform extraction: 

• Add either 700 mL of cold 100% ethanol or 350 mL room temperature isopropanol to the solution to precipitate the plasmid DNA. 
• Add an equal volume of TE-saturated phenol-chloroform to the aqueous DNA sample. 
• Vortex microfuge tube for 30-60 sec. 
• Centrifuge the tube for 5 min at room temperature on the highest setting. 
• Pipette the aqueous DNA layer and place it in a new microfuge tube. 
• Add equal volume of chloroform to the recovered aqueous DNA layer. 
• Repeat steps (ii)-(iv). 

• Add either 700 mL of cold 100% ethanol or 350 mL room temperature isopropanol to the solution to precipitate the plasmid DNA.
• Pour out the supernatant. 
• Wash the pellet with 70% ethanol. 
• Air dry the pellet (can be done by inverting the tube at an angle over Kim wipe) for 20-30 minutes. 
• Resuspend pellet with 25-50 mL of TE. 

Thermofisher MiniPrep Kit Protocol

1. Preheat aliquot of TE to 65°-70°C in water bath. 
2. Secondary culture growth allowed for 14-16 hours- above 16 cell lysis starts. 
3. Pellet and remove supernatant. 
4. Add 250 ml resuspension buffer R3 to resuspend pellet. 
5. Add 250 ml lysis buffer. Mix by inverting till homogenous. DO NOT VORTEX. Incubate at RT for 5 minutes. 
6. Add 350 ml Precipitation buffer N4. Mix by inverting. DO NOT VORTEX. Centrifuge @12000 g for 10 minutes. 
7. Load supernatant into spin column in 2 ml wash tube. Centrifuge @12000 g for 1 minute. Discard flowthrough. 
8. Add 700 ml wash buffer to the spin column. Centrifuge @12000 g for 1 minute. Discard flowthrough, place column back. Centrifuge @12000 g for 1 minute. Discard wash tube. 
9. Place spin column in 1.5mL elution tube. Add 75 mL preheated TE buffer and incubate for 1 minute at RT. 
10. Centrifuge @12000 g for 2 minutes. Discard spin column, elution tube has the plasmid DNA. Store at 4°C. 

Transformation

• Preparation of Competent Cells: Prior to transformation, recipient cells are made “competent” through treatments that increase their permeability. Chemical methods, heat shocks, or electroporation are employed to create temporary openings in the cell membrane, allowing the uptake of foreign DNA.
• DNA Uptake: The exogenous DNA, which can carry genes of interest, regulatory elements, or markers, is mixed with competent cells. The treated cells are exposed to specific conditions, such as a brief electrical pulse or heat shock, that further increase membrane permeability, enabling the DNA to enter the cells.
• Integration or Maintenance: Following uptake, the fate of the exogenous DNA varies. In some cases, the foreign DNA may be integrated into the host genome through homologous recombination. Alternatively, it may persist as an episome, an independent genetic element within the cell.
• Expression of Transgenes: Once integrated or retained, the exogenous DNA can confer new traits upon the transformed cells. This might include antibiotic resistance, production of specific proteins, or alteration of metabolic pathways.

Competency 

1. 500 µL of stationary phase cells subcultured into 50mL LB (1:100 dilution) 
2. Incubate under 37°C till OD reached 0.3. 
3. Incubate cells on ice for 30 minutes. 
4. Pellet down @ 4000 rpm, 4°C(precooled) for 10 minutes. 
5. Discard supernatant.  
6. Add 5 mL 0.2M CaCl₂ and resuspend pellet. 
7. Ice incubation for 30 minutes 
8. Pellet down @ 4000 rpm, 4°C(precooled) for 10 minutes 
9. Add 2mL 0.2M CaCl₂ in 12.5% glycerol.
10. Aliquot 200 mL and store @ -80°C 

Transformation 

1. Add 10mL of Nucleic Acid to 200mL of competent cells and mix gently. 
2. Incubate on ice for 30 minutes. 
3. Heat shock at 42°C for 90 seconds. 
4. Transfer to ice for 5 minutes. 
5. Add 800mL of LB to make total volume 1mL, incubate @ 37°C for 1 hour. 
6. Pellet down at 7000 rpm for 2 minutes, throw the supernatant, and resuspend in the leftover supernatant.
7. Spread on an antibiotic plate.

Modifications:

1. Use 0.1M MgCl₂ instead of 0.2M CaCl₂ in step A6.

Protein Expression

1. Protocol:

• Primary Culture:
  • Add required amount of antibiotic into 5mL LB.
  • Pick a single colony from the transformed plate and add it to LB media containing antibiotic.
  • Incubate at 37°C at 180 rpm overnight
• Secondary Culture
  • Prepare 200mL of antibiotic mixed LB.
  • Add 1% of overnight primary culture into the media: 2mL in this case.
  • Incubate at 37°C at 180 rpm for 3-4 hours.
• Induction:
  • After 3-4 hours measure the OD of the culture.
  • If OD is between 0.6-0.8, add 200µL of IPTG (of 1M working stock solution).
  • Incubate at 25°C at 180 rpm overnight.

Optimisation of Expression:

Our first few experiments had resulted in low expression in protein. To get better yields of induced protein, the optimization of the concentration of IPTG (which induced protein expression) and incubation for expression was conducted as follows:

IPTG Concentration (mM)	Incubation Time (Hours)
1.5	16
2	16
2.5	16
1	24
1	32
1	48
Uninduced Control	NA

These were monitored and a western blot of the cultures were done after all samples were collected.

The last row without signal is the uninduced control sample.

The expression was low in all the samples. This made us go back to choosing the bacterial strain.

Choosing the bacterial strain

Our initial experiments were conducted with the E. coli strain TG1. To improve yields for further downstream applications, we decided to induce expression of antibody in both the TG1 and BL21 strains (both at 1mM IPTG). Then, we ran a western blot to compare the results.

From the results, it was clear that BL21 has a much higher protein production capacity compared to TG1. Hence all further experiments were conducted using BL21 strain of E. coli.

Protein Purification

Training Protocol 

• Lysis of Pelleted E coli
  • 10 ml of lysis buffer is required per gram of pellet. Here we had 4 g but took 50 ml to reduce the viscosity of the solution. We had to add extra components to the lysis buffer not mentioned above. PMSF and lysozyme, PMSF is a protease inhibitor. We made 100 milli molar, but in the final buffer, we need 1 mm.  
  • We pelleted a 500ml E coli culture. 
  • We need 0.5mg lysozyme per mL so for 50 mL we used 25mg.  
  • The adding part was done in a Thermocol ice container with a steel glass, where we mix everything.  
  • Mix the lysozyme into the lysis buffer, be sure to mix multiple times taking a small amount at a time. Take 500 microlitres of PMSF (1 mm) in the 50 ml lysis buffer falcon tube (PMSF is dissolved in alcohol). Vortex well. 
  • Resuspend the pellet solution using a syringe and take 10ml of lysis buffer at a time, resuspend it, and pour in steel glass. At the end you can use a spatula to take remaining pellets. In this process you can use some extra lysis buffer to properly clean it. 
• Ultra Sonification 

This machine produces high-frequency sound waves that help lyse the cell for all the internal matter to come out.!! Important !!- the frequency, pulse time pulse out, and amplitude percentage need to be specific to the cell (E coli) and protein. It shouldn’t degrade the protein.  In this experiment, we used 2 min net pulse time, Pulse on 1s, Pulse out 2s, and Amplitude 50%. Repeat a total of two times with 5 min gap in the middle. 

• In two 50ml falcon tubes, pour the sample avoiding bubbles, balance each one of them to a difference of 0.00g. They must be the same weight or else the centrifuge would make sound and shut down.  
• Centrifuge for 45 minutes at 14000g rpm 
• Setting up the column tube for putting the Ni resin.
  • 1mL resin used for a 20mL column. If we are using soluble proteins; we must collect the supernatant after the centrifugation. Since madam had membrane proteins, she had to collect the resin for the purification.  
  • Collect supernatant and tumble it for one day. 
• Column tube
  • Use a well-washed column tube with a filter at the bottom. Add the Ni resin at the bottom and then add ethanol 10 mL two times, let it drain, and then repeat with water twice. 
  • Then repeat the same wash with lysis buffer, to get the same environment of the column as in the protein-rich supernatant.  
• To confirm the existence of our protein in the supernatant we must run a SDS page both before and after centrifugation. We must know the weight of our protein to detect it using the ladder.  

Note: The term CV in several protocols is a ratio to the amount of resin you are using, not a measure of mL. If we use a resin of 0.5mL and protocol says to add 10CV of some buffer, we add 5mL, as it is relative to the amount of resin we use.  

• Then take the resin and put it in the supernatant falcon tube. Use several to and fro dilutions between the column and falcon tube to get the entire resin in the tube. Avoid bubbles.  
• Tumble it for one hour at 4°C. 
• Then pour the supernatant mixture into the column, all but the resin attached to the protein would be filtered out. Add 10CV of lysis buffer.  

Important: Do not throw away the solution that gets filtered out; we must do a SDS page to check the presence of our protein in it.  

• Add 20CV of wash buffer and collect it, run SDS page. 
• Add elution buffer by dividing it in four parts, if protocol says 4CV, use 1CV at a time, collect the sample and run SDS page for each.
• Then Run dialysis for removing salts.

Ammonium sulphate Precipitation

• A saturated solution of ammonium sulphate was prepared (38.34g in 50ml water). The pH was adjusted to 6.8-7 using 5M NaOH.
• The supernatant was mixed with the saturated solution to make a 70% saturated solution.
• The mixture was kept at 4°C for 6-8 hours/overnight.
• It was then centrifuged at 10,000 rpm for 30 minutes to precipitate the protein.
• The pellet was resuspended in the required buffer.
• The resuspended was then dialyzed overnight to remove excess ammonium sulphate.
• The sample is now ready for purification.

Desalting Buffer

• 1X Tris Cl (pH=8): 50mM
• NaCl: 300mM
• Glycerol: 10%
• MilliQ water

Dissolution of Pellet:

• 1X Tris Cl (pH=8): 50mM
• Glycerol: 10%
• MilliQ water

His-tag purification (using spin kit)

• The spin columns were equilibrated to working temperature. Purifications were performed at 4°C.
• Columns were centrifuged at 700 g for 2 minutes to remove storage buffer. It was then washed with water a couple of times, then equilibrated with the equilibration buffer.
• Columns were centrifuged at 700 g for 2 minutes to remove the equilibration buffer.
• The sample was added to the column and tumbled at 4°C overnight for better binding.
• Columns were centrifuged at 700 g for 2 mins to remove flowthrough.
• 2CV of Wash buffer-1 was added and then centrifuged at 700 g for 2 mins to remove flowthrough. Collect it separately. This was repeated with Wash Buffer-2.
• For elution, 1CV of elution buffer was added each time and centrifuged at 700 g for 2 mins to collect eluent. All eluted fractions were collected separately and labelled accordingly.

The process is analyzed by running a SDS-PAGE with aliquots of sample collected after each step.

Urea extraction

• Spin down cells from large culture at 6000 rpm for 20 min.
• Suspend the cell pellet in 6ml of lysis buffer.
• Sonicate the suspension to lyse all the cells – 45% amplitude, 1min, 1 sec ON, 2s OFF.
• Centrifuge cell lysate for approximately 20 min at 15,000 rpm, 4°C. Discard the supernatant.
• Wash the inclusion body pellets in a small volume of wash buffer(1-2ml). This should solubilize membranes and membrane proteins.
• A short sonication (3 x 10 seconds) is performed during each wash step.
• Centrifuge as above and repeat the washing steps three times.
• Solubilize the purified inclusion bodies into 8M urea with appropriate buffer.
• Leave the IBs to dissolve overnight.
• Run a gel to check for the success of the inclusion body purification.

Lysis Buffer:

• Tris-Cl (pH 8.0): 50mM
• NaCl: 300 mM
• Tween-20: 0.1%
• Triton-X: 1%
• EDTA: 50mM
• DTT: 20mM

Wash Buffer: Lysis Buffer + 2% Triton-X

Resuspending Buffer: Lysis Buffer + 8M urea

Buffer selection for protein purification

To choose the appropriate buffer, the following were considered –

• pI of the protein – for anti-IL8 pI is approximately 5.7.
• Possible buffers:
  • Tris-Cl
  • Phosphate
  • PBS (Phosphate Buffered Saline)
  • HEPES.
• Additives to be added:
  • PMSF (protease inhibitor)
  • β-mercaptoethanol (reducing agent)
  • Glycerol (for stabilizing protein)
• Salt content

The following trial buffers were decided upon –

• Phosphate buffer – pH 8.0
• Tris-Cl buffer – pH 8
• PBS buffer – pH 7.4

All buffers used contained 300mM NaCl and 10% glycerol. The results from the SDS-PAGE are shown below.

<insert SDS-PAGE>

The results from this were inconclusive due to low protein yield.

Later, it was observed that Tris-Cl pH 7.4 buffer was ideal for our protein, as it led to better binding of our protein with IL8 compared to PBS and other such buffers.

Determination of concentration of ammonium sulphate

Due to the low yield from purification, all SDS-PAGE results were inconclusive. To increase protein yield and concentrate the supernatant, we decided to use Ammonium Sulphate precipitation.

2ml samples containing the following % of ammonium sulphate were prepared.

• 40%
• 50%
• 60%
• 65%
• 70%

The pellets were run on an SDS-PAGE to check the result.

https://static.igem.wiki/teams/4829/wiki/pages-photos/engineering/gelammonium-sulp.jpg

70% saturation of ammonium sulphate was chosen as it gave the highest relative yield.

Western Blot

In essence, these blotting technique uses the principles of molecular complementarity (base pairing) to detect and analyze specific molecules (proteins) within complex mixtures, enabling researchers to study gene expression, identify mutations, and analyze protein profiles.

Set up gel for western blot:

1. Prepare the 10% stacking gel solution, assemble the rack for gel solidification.
2. Add stacking gel solution carefully until the level is equal to the green bar holding the glass plates. Add butanol to the top to block air supply. Wait for 15–30 minutes until the gel solidifies.
3. Overlay the stacking gel with the separating gel, after removing the water. (Tip: It is better to tilt the apparatus and use a paper towel to remove the water).
4. Insert the comb, ensuring that there are no air bubbles.
5. Wait until the gel is solidified. (Tip: Solidification can be easily checked by leaving some gel solution in a tube).

Electrophoresis

1. Pour the running buffer into the electrophorator.
2. Place gel inside the electrophorator and connect to a power supply. (Tip: When connecting to the power source always connect red to red, and black to black).
3. Make sure buffer covers the gel completely and remove the comb carefully.
4. Load marker (6 μL) followed by samples (15 μL) into each well.
5. Run the gel with low voltage (60 V) for 6 hours, or until the dye front runs off the bottom of the gel.

Electrotransfer

1. Cut 2 sponge pads and a polyvinylidene fluoride (PDVF) membrane with the same dimensions as the gel. (Pref bigger)
2. Wet the sponge and filter paper in transfer buffer and wet the PDVF membrane in methanol.
3. Separate glass plates and retrieve the gel.
4. Create a transfer sandwich as follows:
   1. Sponge
   2. Gel
   3. PVDF (Ensure there are no air bubbles between the gel and PVDF membrane and squeeze out extra liquid)
5. Relocate the sandwich to the semi dry transfer apparatus. Add transfer buffer to the apparatus and ensure that the sandwich is covered with the buffer. Place electrodes on top of the sandwich, ensuring that the PVDF membrane is between the gel and a positive electrode.
6. Transfer for 90 minutes. (The running time should be proportional to the thickness of the gel, so this may be reduced to 45 minutes for 0.75 mm gels).

Blocking and antibody incubation:

1. Block the membrane with 5% skim milk in TBST for 1 hour.
2. Add primary antibody in 5% bovine serum albumin (BSA) and incubate overnight in 4ºC on a shaker.
3. Wash the membrane with TBST for 5 minutes. Do this 3 times. (Tip: All washing, and antibody incubation steps should be done on a shaker at room temperature to ensure even agitation).
4. Add secondary antibody in 5% skim milk in TBST and incubate for 1 hour.
5. Wash the membrane with TBST for 5 minutes. Do this 3 times.

Visualisation:

1. Wash the membrane with TMB/H₂O₂ and visualise in the gel docker.

Agarose Gel Electrophoresis

Agarose gel electrophoresis is a technique used to separate DNA fragments based on their size and charge. The process occurs within a gel matrix made from agarose; a polysaccharide derived from seaweed. DNA samples are loaded into wells at one end of the gel, and an electric field is applied. The negatively charged DNA molecules move towards the positively charged electrode, propelled by the electric field. The agarose gel acts as a molecular sieve, slowing down the migration of larger DNA fragments more than smaller ones. This creates distinct bands along the gel, each representing DNA fragments of specific sizes. To visualize the separated DNA fragments, a DNA stain is often used. This stain binds to the DNA molecules, making them visible under ultraviolet light. By comparing the migration of unknown DNA samples to that of known DNA markers of known sizes, scientists can accurately determine the size of the fragments.

Preparation of the Gel

1. Weigh out the appropriate mass of agarose into an Erlenmeyer flask. Agarose gels are prepared using a w/v percentage solution. The concentration of agarose in a gel will depend on the sizes of the DNA fragments to be separated, with most gels ranging between 0.5%-2%. The volume of the buffer should not be greater than 1/3 of the capacity of the flask.
2. Add running buffer to the agarose-containing flask. Swirl to mix. The most common gel running buffers are TAE (40 mM Tris-acetate, 1 mM EDTA) and TBE (45 mM Tris-borate, 1 mM EDTA).
3. Melt the agarose/buffer mixture. This is mostly done by heating in a microwave but can also be done over a Bunsen flame. At 30 s intervals, remove the flask and swirl the contents to mix well. Repeat until the agarose has completely dissolved.
4. Add ethidium bromide (EtBr) to a concentration of 0.5 µg/ml. Alternatively, the gel may also be stained after electrophoresis in running buffer containing 0.5 µg/ml EtBr for 15-30 min, followed by destaining in running buffer for an equal length of time.

Note: EtBr is a suspected carcinogen and must be properly disposed of per institution regulations. Gloves should always be worn when handling gels containing EtBr. Alternative dyes for the staining of DNA are available; however, EtBr remains the most popular one due to its sensitivity and cost.

1. Allow the agarose to cool either on the benchtop or by incubation in a 65°C water bath. Failure to do so will warp the gel tray.
2. Place the gel tray into the casting apparatus. Alternatively, one may also tape the open edges of a gel tray to create a mould. Place an appropriate comb into the gel mould to create the wells.
3. Pour the molten agarose into the gel mould. Allow the agarose to set at room temperature. Remove the comb and place the gel in the gel box. Alternatively, the gel can also be wrapped in plastic wrap and stored at 4°C until use.

Setting up of Gel Apparatus and Separation of DNA Fragments

1. Add loading dye to the DNA samples to be separated. Gel loading dye is typically made at 6X concentration (0.25% bromophenol blue, 0.25% xylene cyanol, 30% glycerol). Loading dye helps to track how far your DNA sample has traveled, and allows the sample to sink into the gel.
2. Program the power supply to desired voltage (1-5V/cm between electrodes).
3. Add enough running buffer to cover the surface of the gel. It is important to use the same running buffer as the one used to prepare the gel.
4. Attach the leads of the gel box to the power supply. Turn on the power supply and verify that both gel box and power supply are working.
5. Remove the lid. Slowly and carefully load the DNA sample(s) into the gel. An appropriate DNA size marker should always be loaded along with experimental samples.
6. Replace the lid to the gel box. The cathode (black leads) should be closer the wells than the anode (red leads). Double check that the electrodes are plugged into the correct slots in the power supply.
7. Turn on the power. Run the gel until the dye has migrated to an appropriate distance.

Observing Separated DNA fragments

1. When electrophoresis has completed, turn off the power supply and remove the lid of the gel box.
2. Remove gel from the gel box. Drain off excess buffer from the surface of the gel. Place the gel tray on paper towels to absorb any extra running buffer.
3. Remove the gel from the gel tray and expose the gel to UV light. This is mostly done using a gel documentation system. DNA bands should show up as orange, fluorescent bands. Take a picture of the gel.

SDS PAGE

Gel Preparation

• Clean the glass plates and spacers of the gel casting unit with deionized water and ethanol.
• Assemble the plates with the spacers on a stable, even surface.
• Prepare resolving gel solution using the following volumes (for 10 mL) depending on the percentage of gel required (TEMED should be the last component to be added).

Gel %	Water (mL)	30% acrylamide (mL)	1.5 M Tris-HCl, pH 8.8 (mL)	10% SDS (µL)	10% APS (µL)	TEMED (µL)
8%	4.6	2.6	2.6	100	100	10
10%	3.8	3.4	2.6	100	100	10
12%	3.2	4.0	2.6	100	100	10
15%	2.2	5.0	2.6	100	100	10

• Pour the gel solution in the plates assembled with spacers. To maintain an even and horizontal resolving gel surface, overlay the surface with water or isopropanol.
• Allow the gel to set for about 20-30 min at room temperature.
• Prepare stacking gel solution using the following volumes (for 10 mL): (TEMED is again the last component to be added).

Gel %	Water (mL)	30% acrylamide (mL)	1.5 M Tris-HCl, pH 8.8 (mL)	10% SDS (µL)	10% APS (µL)	TEMED (µL)
5%	5.86	1.34	2.6	100	100	10

• Discard the overlayed water or isopropanol on the resolving gelhttps://www.sigmaaldrich.com/IN/en/product/sigma/E7023
• Add the 5% stacking gel solution until it overflows. Insert the comb immediately ensuring no air bubbles are trapped in the gel or near the wells.
• Allow the gel to set for about 20-30 min at room temperature.

Sample Preparation

• To a volume of protein sample (cell or tissue lysate), add equal volume of loading buffer.
• Boil the above mixture at 95 °C for 5 min. Centrifuge at 16000g for 5 min.
• These samples can be stored at -20 °C or may be used to proceed with gel electrophoresis.

Gel Staining

Coomassie Blue staining: Staining of protein gels with Coomassie Brilliant Blue R-250 is a common procedure to visualize proteins resolved by SDS-PAGE. It is highly sensitive and is suitable for long-term storage of the gels.

• After the electrophoresis, place the gel in a plastic tray containing gel fix solution. Place the tray on a rocking table and fix the proteins for 2 hours.
• Remove the gel fix solution and add Coomassie solution.  Place on a rocking table and stain the gel for 2-4 hours.
• After the staining step, wash the gel several times with distilled water to remove excess stain.
• Add destain solution to the gel. Place on rocking table and destain for about 4 hours till clear blue bands on clear background are visible.
• After destaining, the gels may be stored in gel storage solution and photographed as required.

IVT

In vitro translation, also known as cell-free translation or protein synthesis, is a pivotal molecular biology technique that allows for the synthesis of proteins outside of living cells. This process closely mimics the natural protein synthesis that occurs within cells but takes place in a controlled laboratory setting. The key components of in vitro translation include a cell-free extract, which contains the necessary cellular machinery for transcription and translation, as well as a DNA or RNA template encoding the protein of interest.

The process begins with the preparation of a cell-free extract, typically obtained from a suitable source like E. coli or wheat germ, which contains ribosomes, tRNAs, amino acids, and other essential components for protein synthesis. The extract is then combined with the DNA or RNA template, and protein synthesis is initiated through transcription to produce mRNA and translation to synthesize the target protein.

In vitro translation offers several advantages, including the ability to study specific aspects of protein synthesis, such as translational control or post-translational modifications, in a highly controlled environment. It is also invaluable for producing large quantities of recombinant proteins for various research and biotechnological applications, including drug development and structural biology studies.

We used the ThermoFisher Mega Clear kit and followed the same protocol for this procedure.

Ribogreen Assay

The Ribogreen assay starts with the preparation of a sample containing the nucleic acids of interest. A precisely measured amount of Ribogreen dye is then added to the sample, where it selectively binds to RNA or DNA molecules, inducing a significant increase in fluorescence. The resulting fluorescent signal is directly proportional to the concentration of nucleic acids present in the sample, allowing for accurate and sensitive quantification.

We used the Quant-iT™ RiboGreen™ RNA Reagent and Kit, whose protocol can be found in the protocols section.

Transfection into HeLa cells

Transfection is a fundamental molecular biology technique used to introduce foreign genetic material, such as DNA or RNA, into eukaryotic cells. This process serves as a critical tool in various research areas, including functional genomics, gene therapy, and protein expression studies. Transfection enables scientists to manipulate and study gene expression, investigate ss functions, and develop novel therapies.

Transfection can be achieved through various methods, such as chemical transfection, electroporation, lipofection, and viral vectors. We have used lipofection for our purposes. Lipofection utilizes lipid-based carriers to transport genetic material across the cell membrane.

1. Prepare complexes using a DNA (μg) to Lipofectamine 2000 (μg) ratio of 1:2 to 1:3 for most cell lines.
2. Adherent cells: One day before transfection, plate 0.5-2 x 10⁵ cells in 500 μL of growth medium without antibiotics so that cells will be 70-90% confluent at the time of transfection. Suspension cells: Just prior to preparing complexes, plate 4-8 x 10⁵ cells in 500 μL of growth medium without antibiotics.
3. For each transfection sample, prepare complexes as follows:
   1. Dilute DNA in 50 μL of Opti-MEM I Reduced Serum Medium without serum (or other medium without serum). Mix gently.
   2. Mix Lipofectamine 2000 gently before use, then dilute the appropriate amount in 50µL Opti MEM I medium. Incubate for 5 minutes at room temperature.

Note: Proceed to step (iii) within 25 minutes.

1. After the 5-minute incubation, combine the diluted DNA with diluted Lipofectamine 2000 (total volume = 100 μL). Mix gently and incubate for 20 minutes at room temperature (solution may appear cloudy). 

Note: Complexes are stable for 6 hours at room temperature.

1. Add the 100 μL of complexes to each well containing cells and medium. Mix gently by rocking the plate back and forth.
2. Incubate cells at 37°C in a CO₂ incubator for 18-48 hours prior to testing for transgene expression. Medium may be changed after 4-6 hours.
3. For stable cell lines: Passage cells at a 1:10 (or higher dilution) into fresh growth medium 24 hours after transfection. Add selective medium (if desired) the following day.

LNP formulation

Lipid nanoparticles are intricate assemblies of lipids that exhibit remarkable potential for targeted drug delivery and gene therapy. These nanoparticles are typically composed of phospholipids, cholesterol, and other lipid components, forming a stable, biocompatible vesicle structure. Their unique physicochemical properties enable the encapsulation of hydrophobic and hydrophilic payloads, including small molecule drugs, nucleic acids, and therapeutic proteins.

Lipid nanoparticles excel in overcoming biological barriers, such as the blood-brain barrier or cellular membranes, facilitating precise and efficient cargo delivery. This controlled delivery can enhance therapeutic efficacy, reduce side effects, and enable the use of previously challenging drug candidates.

Protocol for iLNP device preparation

1. Place required materials on a Aluminium foil to keep the area clean.
2. PDMS membrane
   1. In a cup mix 10 parts PDMS and 1 part curing agent (SYLGARD 184)- Mix vigorously for 2-3 minutes.
   2. Remove bubbles in mixture (degassing) using vacuum for 10-20 minutes.
3. Setting Mould
   1. On a silicon wafer having the required design for the device etched onto it, apply an acrylic mould of appropriate size to hold the liquid mixture in place while curing.
   2. Application of the acrylic mould is achieved by using the liquid as an adhesive around the rim, followed by rotation to remove air bubbles.

Note: clean the wafer and the mould in the following order: tissue paper, acetone distilled water and flushing with N₂.

1. Heat at 110°-120°C for 10-15 minutes to allow the mould and wafer to bind.
2. PDMS Pouring
   1. Pour the degassed mixture into the Mould- Wafer bowl from the sides, to avoid bubble formation.
   2. Degas the setup for 30 minutes. Leftover bubbles on the surface can be pushed to the side with a micropipette tip. Care should be taken to prevent bubble formation on the wafer.
   3. Heat the setup at 110°-120°C for 30 minutes for curing, then cool to room temperature in about 10 minutes.
3. Extracting PDMS device
   1. Using a surgical knife make cuts on the inside rim of the mould (without touching the wafer). [Do not scratch the wafer.]
   2. Carefully separate the PDMS from the mould and wafer.
   3. Cover the wafer side of the PDMS with scotch tape to prevent contamination.
   4. As each wafer would have etchings for multiple devices, cut up the PDMS into individual pieces.
   5. Insert a biopsy puncher into the circular marks for input and output for fluids to make holes running across the entire width of the device.
4. Plasma Bonding
   1. Clean a glass slide using the same procedure mentioned before.
   2. Remove tape for the device.
   3. Place in a plasma chamber. Create a vacuum of about 0.35-0.4 psi and run the chamber for about 1-2 minutes.
   4. Take out the glass slide and the device and stick both together such that the taped surface is in contact with the slide.
   5. Apply slight pressure to aid binding.
   6. Heat the slide- device at 110°-120°C for 10 minutes, then cool.

LNP mRNA combination:

• Both the runs were done on the 20-baffle mixer: 2 separate ones were used.
• The flow rate ratio (FRR) was 3:1 (aqueous: lipid).
• The RNA was diluted in citrate buffer to a concentration of 50 µg/ml.
• The lipids were dissolved in ethanol. The ethanol was heated to a temperature of 40°C and the lipids were then dissolved in it.
• 20% w/v solution of sucrose in PBS was prepared.
• The four lipids used were ALC-0315, mal-PEG 2000, cholesterol and DSPC. ALC-0315 was the cationic lipid. It was necessary as RNA is negatively charged and the cationic lipid helps in the uptake of the RNA. The molar ratios used were ionizable cationic lipid: neutral lipid: cholesterol: PEG-ylated lipid:: 50: 10: 38.5: 1.5. DSPC is the neutral lipid.
• The RNA solution was flowed from one side and the lipid solution from the other.
• The final output was collected.
• The flow rate of the RNA was 240 µL per minute and the lipids were flowed at 80µL per minute.
• The LNPs were dialysed after production in 20% PBS-Sucrose.
• In the first run we had obtained close to 500µL of sample. This was used for transfection. In the second run we obtained close to 600µL of the sample (all three runs combined).
• For DLS characterization we diluted according to our need, and we also performed ribogreen assay to check the RNA uptake efficiency.

Aptide conjugation

• Mal-PEG₂₀₀₀-DSPE (2mg) in chloroform (80µL) was added to a solution of cysteinylated APT_EDB (1 mg) in DMSO (80µL) under inert atmospheric conditions.
• The mixture was gently vortexed for 12 hours.
• A speed vacuum concentrator is used to evaporate the solvent.
• The aptide-conjugated phospholipid is characterized sing matrix assisted laser desorption/ionization time of flight mass spectrometry and a Pepti-Gel peptide PAGE analysis Kit.

General Lab Procedures

• LB broth:

Readymade: 25 gm powder/ 1000mL water

Handmade: 1% tryptone + 1% NaCl + 0.5% Yeast extract.

• LB-Agar: 40gm powder/ 1000mL water (Don’t keep at temperatures <50°C)
• SM broth:

1. 1% skimmed milk powder in prepared LB solution.
2. 1% peptone + 1% NaCl + 0.5% Yeast Extract + 1% Skimmed milk powder

Note: DO NOT autoclave SM broth- high temperature causes milk protein polymerization. Rather, follow 1, take autoclaved LB and add milk powder at about 60°C.

• Autoclaving:

1. Loosen caps of falcons, Eppendorf tubes and jars and cover jars with Aluminum foils to prevent evaporation loss.
2. Release pressure completely before opening the hatch.
3. Use autoclave gloves and lab coats while retrieving stuff.
4. When putting falcons and agar plates inside, put them in autoclavable bags and loop the open end before rubber banding them, to seal the water vapor out.

Experimental Ca²⁺ based assay

• This assay was based on the fact that a characteristic Ca²⁺ release from the ER within the cell is expected after the activation of CXCR2 in macrophages and neutrophils. The protocol we used is as follows:

1. We seeded a 96 well plate with RAW cells as follows: Media without antibody Media without antibody Media without antibody Media with antibody Media with antibody Media with antibody Media with Antibody+IL8 Media with Antibody+IL8 Media with Antibody+IL8 Media with IL8 Media with IL8 Media with IL8 No dye No dye No dye
2. We used the required amount of FLUO-4AM, (4 μM) in the wells, incubated for 30 minutes.
3. Then, we threw out the old media washed with PBS, added our media’s (as shown above). Incubate for 13 minutes.
4. Finally, we removed this media, added the following buffer: 136 mM NaCl, 4.8 mM KC1, 1 mM CaC12, 5 mM glucose, and 20 mM HEPES, adjusted to pH 7.4 and 310 mOsm.
5. Use a microplate reader to read fluorescence: 494nM excitation, 510nM emission.

This protocol is based off the following papers: [28, 29]