1.Plasmids Midi-Purification
Preparation
The GeneAll plasmid DNA extraction kit was used for the plasmid DNA extraction. For first extraction, 160 ml of absolute ethanol (ACS grade or better) was added into Buffer EW2 as indicated on the bottle. Add RNase A to Buffer P1 and store it at 4˚C. Chill Buffer P3 for better result. Buffer P3 can be stored at 2~8˚C without any precipitation. Buffer P2 and EG may precipitate at cold ambient conditions. If precipitate appears, dissolve it in the 37˚C water bath.
Prepare ice. Prepare the 37˚C water bath or incubator. All experiments should be performed at room temperature.
Unless specified differently, centrifugation should occur at room temperature using a standard centrifuge with the capability to achieve 4,000 to 5,000 x g. Ensure the centrifuge is equipped with a swinging-bucket rotor.
Procedure
Preparation of Cleared Lysate
1.Pellet 50~150 ml of bacterial culture by centrifugation for 5 min at 10,000 x g in a tabletop centrifuge. Discard the supernatant as much as possible.
• Use the appropriate amount of bacterial for better yield of DNA: up to 150 ml of a bacterial culture can be used when the A600 is less than 1.8. For the bacterial culture with high cell density (A600>2.0), reduce the starting sample volume to 50 ml. Too high cell mass of the starting sample can cause the reduction of lysis efficiency and the clogging of the columns, resulting in unsatisfactory yields.
• Bacterial cultures were for 16 to 24 hours in LB-broth containing selective antibiotics.
2.Resuspend pelleted bacterial cells thoroughly in 4 ml of Buffer P1. It is essential to thoroughly resuspend the cell pellet.
3.Add 4 ml of Buffer P2 and mix by inverting the tube 5~6 times (DO NOT VORTEX). Incubate until the cell suspension becomes clear and viscous, but DO NOT incubate for more than 5 min.
• It is important to proceed to the next step immediately after the lysate becomes clear without any cloudy clumps.
• If precipitated material has formed in the bottle of Buffer P2, heat to dissolve at 37˚C (or above). Use of precipitated Buffer P2 may cause significant decrease in DNA recovery yield.
•Keep Buffer P2 capped tightly after use because it can be acidified gradually when in contact with air
4.Add 4 ml of Buffer P3 and thoroughly but gently mix by inverting the tube 7~8 times (DO NOT VORTEX).
• Mix the solution gently but completely and immediately after addition of Buffer P3 for optimal precipitation.
• Use of pre-chilled Buffer P3 or incubation on ice may help the precipitation and the removal of the cellular debris more efficiently, and it may reduce the possibility of the contamination of chromosomal DNA.
5. Pour all of the lysate into EzClearTM Filter (blue ring) sitting on a 50 ml conical collection tube. Let it stand for 2 min and centrifuge for 2 min at 1,500 x g (2,800 rpm).
• Cellular debris will rise to the top during incubation, and this will assist the clearing of lysate through EzClearTM Filter. Failure to perform the incubation may lead to incomplete filtration of lysate. A small amount of liquid can remain trapped in the residual insoluble material, but this will not lead to noteworthy decrease in yield.
• If the cell mass of the bacterial culture is very dense (A600>2.0) and the starting volume is larger than 100 ml, it may be necessary to centrifuge the lysate before transferring to EzClearTM Filter. Because too high cell mass can cause the clogging of the filter on the next step.
• This additional centrifugation can be done at 4,500 x g for 20 min on a swinging-bucket rotor or at 10,000 x g for 10 min on a fixed-angle rotor.
• After this optional centrifugation, transfer the supernatant into EzClearTM Filter unit (Some debris can be co-transferred).
6.Apply 500 μl of Buffer ER to the filtrate and close the cap of the 50 ml conical tube. The volume of filtrate may be about 10 ml.
7.Vortex the mixture and incubate for 15 min on ice.
• The mixture will be turbid with vortexing and then become clear during incubation on ice.
8. Incubate for 15 min at 37˚C and centrifuge for 2 min at 1,500 x g (2,800 rpm).
• After centrifugation, the mixture will be separated by two phases: the white upper phase and the blue (greenish) lower phase. Handle the tube gently not to mixed-up these phases.
9.Transfer carefully the upper phase (clear) into a new 15 ml conical tube (provided) by pipetting.
• Be careful not to co-transfer the lower phase (greenish blue) since it is liable to be dispersed
• If phases are mixed, repeat the centrifugation again.
Purification of Plasmid DNA
10.Add 1/2 volume of Buffer EG to the transfer and invert several times to mix completely.
• For example, 10 ml of the solution, 5 ml of Buffer EG should be added.
11.Transfer all of the mixture to the SV Midi column (clear ring) by decanting or pipetting. Centrifuge for 2 min at 1,500 x g (2,800 rpm). Remove the column, discard the pass-through, and re-insert the column to the collection tube.
12.Apply 10 ml of Buffer EW1 and centrifuge for 2 min at 1,500 x g (2,800 rpm). Remove the column, discard the pass-through, and re-insert the column to the collection tube.
13.Apply 10 ml of Buffer EW2 and centrifuge for 2 min at 1,500 x g (2,800 rpm). Remove the column, discard the pass-through, and re-insert the column to the collection tube.
14.Apply 3 ml of Buffer EW2 and centrifuge for 15 min at 4,500 x g (5,000 rpm). Transfer the column to a new 50 ml conical tube (provided).
• These EW1 and EW2 washing step will remove any traces of lipopolyssacharides (endotoxin), endonucleses, proteins, carbohydrates, and other cellular components bound non-specifically to the column membrane.
• Residual ethanol in eluate may interfere with the subsequent reactions. If carryover of ethanol occurs, incubate the column for 10~20 min at room temperature to evaporate residual ethanol.
15.Add 0.6 ml of Buffer EF or endotoxin-free water directly onto the center of the column membrane and close the cap. Incubate for 5 min at room temperature and centrifuge for 5 min at 4,500 x g (5,000 rpm).
• Ensure that the elution buffer is dispensed directly onto the center of column membrane for optimal elution of DNA.
• The volume of eluent can be increased or decreased as necessity. Higher volume will decrease the concentration of the eluate but yield slightly more DNA.
• The volume can be decreased but it should be 0.5 ml at least, because too small volume of EF buffer may be insufficient to soak the entire membrane.
• For long-term storage of DNA, other buffers, such as TE (1 mM EDTA, 10 mM TrisCl, pH 8.0), can be used for elution. The buffer for elution should be neutral pH (7.0<pH<9.0) and low-salt condition.
• Ensure that the solution and the plastic-ware for elution is endotoxin-free.
16.(optional)
• For higher concentration of eluate; re-load the eluate from step 15 into the column membrane, close the cap, incubate 5 min at room temperature, and centrifuge for 5 min at 4,500 x g (5,000 rpm).
• Increase the total yield; add 0.6~1 ml of fresh Buffer EF into the column, close the cap, incubate 5 min at room temperature, and centrifuge for 5 min at 4,500 x g (5,000 rpm).
The first and second eluates can be combined or collected separately as necessary.
2.Medium Preparation
Dulbecco’s Modified Eagle’s Medium (DMEM) or RMPI medium – 1 liter
• Prepare 900 ml of distilled water in the clean glass beaker. Water temperature should be 15-30ºC. Put the beaker on a stirrer and add a magnet.
• Add the 2 packages of DMEM (ThermoFisher) or Roswell Park Memorial Institute (RPMI) 1640 powder (ThermoFisher) powder to the water and stir gently. Fill some distilled water into the empty package, stir and pour the remains into the beaker. Stir till completely dissolved.
• Add 3.7 gram Sodium Bicarbonate (or 49.4 ml of 7.5% Sodium Bicarbonate Solution; Sigma).
• Adjust pH value to about 7.4 using 1N HCl or 1N NaOH. The pH will rise by 0.1-0.3 units after filtration.
• Top up the volume by Milli-Q water to 1 liter.
• Filter for sterility with 0.2 µm filter (Nalgene) into sterile bottles.
• Add 100 ml heat-inactivated fetal bovine serum (FBS) and 100 U/mL of penicillin and 100 μg/mL streptomycin (Thermofisher).
3.HEK293T Cell Passage
Materials and Reagents
• HEK293T Media (DMEM with 10% FBS with P/S)
• Phosphate Buffered Saline (PBS)
• Trypsin/EDTA (0.25%) solution
• Class 2 Biosafety Cabinet
• 10 and 5 mL sterile pipettes
• Vacuum apparatus w/ autoclaved Pasteur pipets
• T-75 cell culture flask
Steps
1.Prepare Biosafety Cabinet and materials
a.Using UV to sterile the BSC for 15 mins. Turn off the UV light before use.
Turn on venting and lighting.
b.Turn on venting and lighting.
c.Spray down inside surfaces of the cabinet with 70% ethanol and wipe down.
d.Spray with 70% ethanol and wipe down all surfaces before placing them in the cabinet.
2.Use vacuum apparatus and a Pasteur pipet to aspirate the old media.
3.Rinse the cells with 10 mL PBS.
a.Add the medium onto the side of the dish/flask rather than directly on the surface that the cells are laying on to avoid power-washing them away.
5.Aspirate PBS.
a.This time you actually do want to apply the trypsin to the cell-containing surface.
6.Incubate for 3-5 minutes, until all the cells are loosely floating (can be observed under microscope).
7.Add 4 mL complete medium (i.e. with FBS and P/S) and mix until cells are consistently spread through the solution.
8.Transfer all the mix solution into a 15 ml centrifuge tube and centrifuge for 3 minutes with 1000 x rpm and 37 celsius degree.
9.Aspirate the supernatant away and resuspend the cells by 5 ml complete medium.
10.Pipet a calculated aliquot of cell suspension (generally 5-10% of the total) into the new 75 cm^2 cell culture flask and the 100 mm cell culture dish.
11.Add 10 mL complete media to the flask and dish.
12.Incubate the cells in the humidified incubator set to 37oC and 5% CO2.
13.Clean up the Biosafety Cabinet and return reagents/media to the fridge.
4.HEK293T Cell Frozen Stock Preparation
Materials and Reagents
• HEK293T Media (DMEM with 10% FBS and 1% p.s.)
• Phosphate Buffered Saline (PBS)
• Trypsin/EDTA
• Class 2 Biosafety Cabinet
• 10 and 5 mL sterile pipettes
• Vacuum apparatus w/ autoclaved Pasteur pipets.
• 75 cm^2 cell culture flask
• DMSO
• 2 mL freezing tubes
Steps
1.Follow steps 1-7 in the aforementioned procedure.
2.Prepare freezing media.
a.5% DMSO: mix 10 mL complete media with 0.5 mL DMSO.
3.Filter freezing media.
a.Sterile the 5% DMSO solution through a 0.2 mm syringe filter.
b.Do not retract plunger after filtering while the filter is still attached, as this will pull impurities back into the syringe. First disconnect the filter, then retract the plunger, then reconnect the syringe and filter.
4.Chill freezing medium for 3-5 minutes on ice.
5.Centrifuge remaining cells after the passage.
6.Remove the supernatant and resuspend the cells by the chilled freeze medium.
7.Stepwise to reduce the temperature:
a. Place freezing tubes in the Freezing Container -20°C
b. Store the cells at -80°C for 16-24 hours.
c. Finally store the cells in the liquid nitrogen tanks.
5.Transfection with HEK293T Cells (Lentivirus Packaging)
Workflow Timeline
Day 0: Seed the HEK293T cells
Day 1 (pm): Transfect plasmid DNA into the HEK293T cells
Day 2 (am): 18 h post-transfection. Remove media, replace with fresh media.
Day 3-4 (am): Harvest virus
Equipment
• Class II, Type A2 Biological Safety Cabinet
• 0.5–10 µL single channel pipette
• 2–20 µL single channel pipette
• 20–200 µL single channel pipette
• 200–1000 µL single channel pipette
• Ice bucket
CO2 incubator
Pipet controller
Hazardous waste container
pH meter
Stir plate
Magnetic Stir Bar
Reagents
DMEM high glucose
L-alanyl-L-glutamine (or alternative stable glutamine such as glutaGRO, Corning 25-015-CI)
Transfection medium: Opti-MEM or Opti-Pro SFM
25 mM chloroquine
Polyethylenimine (PEI), linear MW 25,000 Da
Heat-inactivated FBS
1x PBS pH 7.4 without calcium or magnesium (cations can affect the attachment of adherent cells)
0.45 μm polyethersulfone filter
Microcentrifuge tubes, Neptune 3745.X
10 cm tissue culture dish
15 mL conical tubes
Hydrochloric acid
Sodium hydroxide
0.22 μm polyethersulfone (PES) filter
Transfection grade DNAs
Procedure
1.Seed HEK293T packaging cells at 3.8×106 cells per plate with DMEM medium in 10 cm tissue culture plates.
2.Incubate the cells at 37 °C, 5% CO2 for ~20 h
3.Prepare a mixture of the 3 transfection plasmids and PEI with the OptiPro SFM (ThermoFisher) medium:
4.Make a mixture of a total of 500 μL PEI-OptiPro SFM with enough PEI such that the ratio of μg DNA:μg PEI is 1:3 (1000 μL total per 10 cm dish).
5.Gently add the diluted PEI mixture to the diluted DNA mixture. Add the diluted PEI dropwise while gently flicking the diluted DNA tube. Incubate the mixture 12–15 min at RT.
6.During the incubation, add 10 mL of DMEM Complete to a 15 mL conical and an appropriate amount of 25 mM chloroquine to the conical so that the final concentration in the dish (once transfection mixes are added) will be 25 uM of chloroquine.
7. After the incubation, add the DNA:PEI-Max mix to the conical containing the media and chloroquine and mix well.
8.Gently aspirate the media out of the previously seeded 10 cm plate.
9.Slowly pipette the transfection mix onto the 10 cm plate, being careful not to disturb the cells.
10.Incubate the cells for 18 h, or until the following morning.
11.The following morning, carefully aspirate the media. Replace the media with 10 mL of DMEM Complete or OptiPro SFM.
12.Incubate the cells in the CO2 incubator.
13.Virus can be harvested at 48, 72, and 96 h post transfection in individual harvests or a combined harvest where all the individual harvests are pooled. If pooling harvests, transfer the harvested media to a polypropylene storage tube and store at 4 °C between harvest.
14.Centrifuge the viral supernatant at 2100 rcf for 5 minutes to pellet any packaging cells that were collected during harvesting.
15.Filter supernatant through a 0.45 μm PES filter.
16.The viral supernatant can be stored at 4 °C for several hours but should be aliquoted and snap frozen in liquid nitrogen and stored at -80 °C as soon as possible to avoid the loss of titer.
6.Infection with HEK293T Cells and Generating Stable Cell Lines with Target Proteins
Workflow Timeline
Day 0: Seed Cells and infection with the lentivirus
Day 2–3 (am): Remove media, replace with fresh media containing selection reagent
Day 3–14: Change medium as needed
Day 14–18: Expand and harvest stable cell lines
Equipment
• Class II, Type A2 Biological Safety Cabinet
• 0.5–10 µL single channel pipette
• 2–20 µL single channel pipette
• 20–200 µL single channel pipette
• 200–1000 µL single channel pipette
• Ice bucket
• CO2 incubator
• Pipet controller
• Hazardous waste container
Reagents
• DMEM high glucose
• L-alanyl-L-glutamine (or alternative stable glutamine such as glutaGRO)
• Heat-inactivated FBS
• Polybrene (10 mg/mL)
• 1X PBS pH 7.4 without calcium or magnesium (cations can affect the attachment of adherent cells)
• 0.45 μm polyethersulfone filter (for viral preps, if prep was not previously filtered)
Microcentrifuge tubes
• Microcentrifuge tubes
• 6-well tissue culture treated dish
• 15 mL conical tubes
• Lentivirus preparation (in experiment 5 above)
• Appropriate antibiotic for gene selection (e.g. puromycin, blasticidin)
Procedure
1.Before beginning, determine the optimal dosage of the antibiotic for your target cell line. To do this, treat target cells with a range of doses of antibiotic and determine the lowest dose that kills all of the cells.
2.Prepare a batch of DMEM complete + 10 µg/mL polybrene by diluting 20 µL of 10 mg/mL polybrene into 20 mL media.
3.Thaw the lentiviral aliquot on ice prior to use.
4.Add 0.5 mL of a single viral dilution to each well (each well gets one dilution, so a 6-well plate will hold 5 dilutions plus one 'no virus' control well).
5.Incubate the cells with the virus for 48–72 h.
6.Gently aspirate the media from the cells.
7.Add 1.5 mL of DMEM complete containing the appropriate antibiotic. This is the beginning of the selection process, which will begin the selection of a stable cell pool.
8.Observe the dish every day to ensure that the cells in the control well (0 µL lentivirus, above) are dying. Perform regular media changes and monitor the growth of the cells.
9.As polyclonal populations of resistant cells start proliferating and the individual wells become confluent, expand into larger vessels. A confluent well of a 6-well dish can be expanded into a 10 cm dish. A confluent 10 cm dish can be expanded into two 75 cm2 flasks, etc.
10.Once the polyclonal populations are growing well and have been sufficiently expanded, prepare cell stocks and/or harvest to test for protein expression.
Typically, cells transduced with lower dilutions of the virus will have higher levels of expression. Consider expanding populations transduced with a variety of dilutions and pick the population that has the most desirable level of expression.
11.Over time, transgene expression in a polyclonal population may drop. This is because cells that express high levels of the transgene may have reduced growth rates, especially if that transgene is toxic. Eventually, the rapidly growing low-level transgene expressors may take over the culture. To overcome this, consider generating monoclonal lines from the early polyclonal populations.
7.Cell Cytometry
Preparation of Adherent Tissue Culture Cell Lines
1.Harvest cells by 0.25% trypsin, followed by quenching with media containing serum.
(Epitopes may be cleaved when using the enzymatic digestion method. Cells can also be harvested by gently scraping them into culture media)
• Remove the culture medium. Eliminate residual serum by rinsing cell monolayers with PBS.
• Slowly add or 0.25% Trypsin to cover the cell monolayer.
• Incubate at 37°C for up to 10 minutes.
• After incubation, gently tap the flask and the cells will detach and slide off in one sheet to the bottom of the flask.
• Add growth medium and re-suspend the cells by gently pipetting.
2.Centrifuge at 400 g for 5 min.
3.Discard supernatant and resuspend pellet in fresh PBS-BSA to wash off any remaining cell debris and proteins.
4.Centrifuge at 400 g for 5 minutes at RT.
5.Discard supernatant and resuspend pellet in an appropriate amount of PBS-BSA.
6.Count cells using a hemocytometer or an automated cell counter. Dilute the cells with cold (4°C) PBS-BSA to a minimum concentration of 1 x 107 cells/mL.
7.The cells with target protein expression was sorted by flow cytometry (BD FACSCalibur)
Flow Cytometry Works
The sample is injected into the center of the sheath stream of flow cytometer in a liquid state; therefore the particles are distributed randomly. The fluidics system is then responsible for separating out the particles into an ordered stream of single particles.
After hydrodynamic focusing, the cells or particles of interest pass through the laser beam therefore intercepting and scattering the light which excites the fluorochromes to a higher energy state. The energy is then released as a photon of light with spectral properties unique to specific fluorochromes. Light scattered in the forward direction (as shown in the below diagram) is collected by a lens which is in line with the beam known as the forward scatter channel (FSC). The FSC intensity gives the particles size and can give information used to distinguish between cellular debris and living cells. The side scatter channel (SSC) is perpendicular to the beam and provides information about the granular content within a particle. Both FSC and SSC are unique for each particle and a combination of the two may be used to differentiate between different cell types in a heterogeneous sample.
Cell Sorting
Fluorescence-activated cell sorting (FACS) is a specialized type of flow cytometry. The rate of flow sorting at 10 000 cells/second provides a method for sorting a heterogeneous mixture of biological cells into separate storage containers. It is based upon the specific light scattering and fluorescent characteristics of each cell. It is an extremely useful scientific instrument, as it provides fast, objective and quantitative recording of fluorescent signals from individual cells as well as physical separation of cells of particular interest.
Cell Sorting Works
After the cells have passed through the laser beam and the detectors, a vibrating mechanism causes the stream of cells to break into individual droplets. An electrical charge is placed at the point where the stream breaks into droplets immediately prior to the fluorescence intensity measurement, and the opposite charge is trapped on the droplet as it breaks from the stream. The droplets then travel through a strong electrostatic field and are deflected based on their charge into waiting sample tubes. The number of cells and level of fluorescence in each tube is then known.
8.PCR
Materials
• DNA or cDNA sample
• 0.50-ml microcentrifuge tube
• P20 micropipette (pipetman)
• Box of micropipette tips (200 mL, yellow)
• Beaker of ice
• Reverse primer and forward primer (Note: these can be in many different concentrations) 0.1 μM to 1 μM
• Mixture of 4 mM each of 4 deoxynucleoside triphosphates
• 10X PCR buffer (1X = 1.5 mM MgCl2, 50 mM KCl, 10 mM TrisHCl, pH 9.0, 1% Triton X-100)
• 1 unit/μl Taq DNA polymerase
• Mineral oil (if thermocycler does not have hot lid to reduce condensation)
• Thermocycler
Polymerase Chain Reaction
• 1.Usually 20 to 50 μl total in volume and will include the following:
• 0.1 to 1 μg of genomic DNA or cDNA, ~0.1μg should be sufficient for plasmid DNA
• 10X PCR buffer to give a final concentration of 1X
• 4 mM dNTP mix (dCTP, dATP, dGTP, dTTP) to give a final concentration of 0.2 mM
• Both the forward and reverse primer added at a final concentration of 0.1 μM to 1 μM of each primer
• 1 unit/μl Taq polymerase
• H2O (DNA and DNase free) to bring volume to 20 μl to 50 μl
An example 20 μl reaction:
• 1 μl of dsDNA template (~0.1 μg)
• 2 μl of 10X buffer 1 μl of 4 mM dNTP mix
• 1 μl of 10 μM forward primer to a final concentration of 0.5 μM
• 1 μl of 10 μM reverse primer to a final concentration of 0.5 μM
• 1 μl of 1 unit/μl Taq polymerase
• 13 μl of water
2. Combine the reagents in the 0.5-ml tube or the 0.2-ml PCR tube. Be sure to keep the reagents on ice.
3. Tap the tube gently to mix and spin briefly in the microcentrifuge to get all contents to bottom
4. Place on ice until ready to load in the thermocycler.
5. If the thermocycler does not have a heated lid, layer thin film of mineral oil over the mixture to prevent evaporation during cycling.
6. Upon completion of PCR, hold samples at 4℃. Prepare the DNA for loading by addition of 1/10 volume stop-loading buffer (contains EDTA, glycerol, and bromophenol blue)
7. Analyze by gel electrophoresis and be sure to include size markers in at least one well on the same gel.
Example Typical Thermal Cycler Program
Step 1: 92 to 98℃, 30 seconds to 1 minute
Step 2: optimal annealing temperature of primers, 37 to 65℃, 30 seconds to 1 minute
Step 3: 72℃, 30 seconds to 1 minute
Repeat steps 1 to 3 for 20 to 30 times to accumulate enough amplified target DNA to be visualized on a gel.
Step 4: 4℃ holding of sample until analysis by gel electrophoresis
Agarose Gel Electrophoresis: UltraPure Agarose
Ingredients
•1 gel casting tray and tape
•1 10-well comb
•1 electrophoresis chamber
•1 500 mL flask
•1 g UltraPure Agarose
•1 L UltraPure 1X TBE
•10 mg/mL ethidium bromide
•1 TrackIt Kb Plus DNA ladder
•1 PCR DNA sample
•1 10X BlueJuice Gel Loading Buffer
Procedures
1.Weigh out 1 g of agarose gel and add it to a 500 mL flask. Add 100 mL of 1X TBE. (The total gel volume will vary depending on the size of the casting tray.)
2.Gently mix and heat the solution in a microwave until the agarose is completely dissolved. (CAUTION: wear protective gloves when handling extremely hot agarose solution and heat the solution in several short intervals—do not let the solution boil for long periods as it may boil out of the flask or cause a loss to water vapor. You can weigh the flask before and after heating and add distilled water to make up for the lost volume.)
3.Allow hot agarose to cool in a water bath set at 50–55°C for 10 min. Swirl the flask occasionally to cool evenly.
4.Prepare the gel casting tray by sealing the ends of the gel chamber with tape. Place the 10-well comb in the gel tray. (Note: pour the gel on a level surface; otherwise, the gel thickness will not be even, resulting in uneven migration)
5.Add 5 µL of stock ethidium bromide solution to the cooled gel and pour into the gel tray. Remove any bubbles and allow the gel to cool for 30 min at room temperature. (Exercise caution when using ethidium bromide, which is a known carcinogen.)
6.Remove the comb and tape, place the gel tray in the electrophoresis chamber, and cover with 1X TBE buffer. (Cover the gel with ~2–3 mm of buffer for best results.)
7.Add 0.2 uL volume of loading buffer to samples; e.g.,4uL of 20 uLsample. Load 100-250 ng ofyoursample and add E-Gel loading buffer to 20 uL.
8.Load the first and last wells with 20 µL of diluted markers/ladders (Use 5 µL of E-Gel marker plus 15 µL of water). Continue loading 20 µL of sample per well.
9.Electrophorese at 100V for 40 min. Make sure that the current is running in the correct direction.
10.After the run, remove the gel with the tray and visualize PCR fragment bands using the E-Gel Imager (Invitrogen) or other blue light source.
9.Exosomes Extraction
Shipping and Storage
The exoEasy Maxi Kit (QIAGEN, cat. no. 76064) is shipped at ambient temperature. Store all components dry at room temperature (15–25°C). All kit components are stable for at least 9 months upon arrival under these conditions.
To ensure compatibility with biological applications, Buffer XE is produced sterile without added preservative to prevent microbial growth. Handling under sterile conditions is recommended. Buffer aliquots can also be stored frozen to prevent growth.
Principle
The exoEasy Maxi Kit uses a membrane-based affinity binding step to isolate exosomes and other EVs from serum and plasma or cell culture supernatants. The method does not distinguish EVs by size or cellular origin and is not dependent on the presence of a particular epitope. Instead, it makes use of a generic, biochemical feature of vesicles to recover the entire spectrum of extracellular vesicles present in a sample. It is therefore essential to completely remove cells, cell fragments, apoptotic bodies, etc., by centrifugation or filtration of samples before starting the protocol.
Particulate matter other than vesicles, like larger protein complexes that are especially abundant in plasma and serum, is largely removed during the binding and ensuing wash step.
After washing the column membrane, intact vesicles are eluted in aqueous buffer containing primarily inorganic salts and are then ready to use for physical or biochemical analysis. For certain applications, such as biological assays, an additional concentrating or buffer exchange step may be required. This can be achieved using ultrafiltration, using a pore size of 100 kDa or smaller.
Important Notes
1.The volume of starting material is limited by the binding capacity of the exoEasy spin column. For cell culture supernatants, up to 32 mL (equals 4 column loading steps) have been processed with good results. However, the concentration of vesicles in supernatants depends strongly on the cell type and culture conditions, and therefore we recommend starting with no more than 16 mL of sample for material that has not been previously tested with the kit. Higher sample volumes may result in reduced recovery of vesicles. It is strongly recommended to only use pre-filtered sample material, excluding particles larger than 0.8 µm. Filtering should be performed prior to freezing samples for storage, if possible.
2. The provided amount of Buffer XBP is sufficient for processing of 20 x 8 mL samples. To process large volumes, additional Buffer XBP can be ordered separately.
3.Vesicle yields can be estimated by different particle analysis technologies such as NTA and TRPS, or by quantitation of vesicular markers (e.g., proteins such as CD63 and RNA).
4.If the recommended sample volume is exceeded, recovery of vesicles will not be consistent and may be reduced.
5.After collection and centrifugation, cell culture supernatant can be stored at 2–8°C for up to 6 h or used directly in the procedure. For long-term storage, freezing in aliquots at –15°C to –30°C or –65°C to –90°C is recommended. To process frozen samples, incubate at 37ºC in a water bath until samples are completely thawed. Avoid prolonged incubation as this may compromise integrity of EVs.
6.All steps should be performed at room temperature (15–25°C). Carry out the protocol steps quickly but carefully.
7.Centrifugation of the exoEasy Maxi spin columns is performed in a standard laboratory centrifuge with a swinging bucket rotor, preferably capable of up to 5000 x g (it is possible to reduce the steps performed at 5000 x g down to a minimum force of 3000 x g without performance loss).
Procedure
1.Supernatants should be filtered to exclude particles larger than 0.8 µm (e.g., using Sartorius Minisart NML (cat. no. 16592) or Millipore Millex-AA (cat. no. SLAA033SB) syringe filters).
2.Add 1 volume buffer XBP to 1 volume of sample. Mix well by gently inverting the tube 5 times. Let the mixture warm up to room temperature.
3.
Add up to 16 mL of the sample/XBP mix onto the exoEasy spin column and centrifuge at 500 x g for 1 min. Discard the flow-through and place the column back into the same collection tube.
If the sample volume is greater than 8 mL, repeat this step until the entire volume has been passed through the column.
Optional: To remove residual liquid from the membrane, centrifuge at 5000 x g for 1 min.
4.Add 10 mL buffer XWP and centrifuge at 5000 x g for 5 min to remove residual buffer from the column. Discard the flow-through together with the collection tube.
Note: It is possible to reduce the centrifugation speed from 5000 x g down to a minimum force of 3000 x g without loss of performance.
Optional: Repeat step 4 to further reduce nonspecifically bound materials (e.g., free proteins). Reuse the collection tube from step 4.
5.Transfer the spin column to a fresh collection tube.
6.Add 250–400 µL Buffer XE to the membrane and incubate for 1 min. Centrifuge at 500 x g for 5 min to collect the eluate.
Optional: Re-apply the eluate to the exoEasy spin column membrane and incubate for 1 min. Centrifuge at 5000 x g for 5 min to collect the eluate.
10.Dynamic Light Scattering (DLS) Measurement
Preparation of Instrument System
1. Switch on the host and the temperature controlled sample unit. Set the desired temperature (range 4-60 ℃). Give the laser at least 20 min to warm up.
2. Start the software (Dynamics V6.3.40).
3. The Dynamics window opens.
4. Create a new experiment (or load an old one if you have one with the right parameters).
5. Check the parameters:
You can change AcqTime, Laser Power and Set Temp here. Good start values are Acq Time 10, Laser Power 10, and Set Temp (C) 21, they can be changed as required later.
Under Solvent:
Viscosity should change as you choose your buffer conditions. If your conditions are not included, you should add them to the list.
Under Sample:
Choose the model you will use here (Globular Protein for protein work), and your concentration Make sure the volume is set to .012mL (12ul). If you want to calculate molar mass enter your dn/dC (dn/dC (mL/g) = 0.187 is right for protein).
6. Click the connect to the instrument button.
7. The round button next to it should turn to green.
8. The instrument should now ramp up to the set temperature.
9. Check the readings coming in by clicking the Instrument Control Panel. A new window opens displaying the incoming readings. You can use this window to control laser power and acquisition time. Without a cuvette in place, readings should be 20,000 cts/s with the laser power at 10%. If they are higher, there is a glitch in the system. Reselect the laser power by clicking on the slider and set the laser power to 10% again.
Preparation of Samples
10. A protein concentration of 1 – 2 mg/ml is a good starting range. For smaller proteins, better readings are obtained with higher concentrations.
11. It is extremely important that your samples are free of any particulates like dust, free of unwanted aggregates, and free of bubbles.
12. Filter your samples, buffer, 100% EtOH, and Milli-Q water through 0.1 µm filters (spin filters for small sample volumes, syringe filters for intermediate volumes of filter discs or for filtering large volumes of buffer). Also make sure that all your sample vials and buffer bottles are dust free.
13. When filtering is not possible due to adverse effects of the filter on the sample (shear forces and binding problems) spin the sample for 15 min in an ultra-centrifuge and only take the top third.
14. To avoid the formation of air bubbles, degas your buffer and give the samples time to reach the desired temperature. This is especially important for samples on ice or samples to be tested at temperatures higher than room temperature.
Buffer Control and Water Control
15. Prepare the cuvette. Again, the cuvette must be clean and dust free. Shared cuvettes should be stored clean, dry and closed on the shelf above the DLS. Don't rely on them being clean. You will measure a water and buffer baseline before you measure your sample. To be sure that the cuvettes are clean and dust free you can rinse them first with filtered water (3 times), then with absolute EtOH (3 times). Dry them by holding them upside down and gently blowing into them using compressed air from a can. Do not use house compressed air; it can contain oil/dust from the compressor and will contaminate the cuvettes strongly. Once the cuvette is dried keep it closed to prevent dust from forming.
16. If you use a 12 µl cuvette fill it with your filtered H2O, using a long gel loading tip. Do not use glass transfer pipettes or steel needles to avoid scratching the quartz windows of the cuvette. If you use a syringe filter system be extremely careful. Use no more than 14 µl and go slow observing through the window to avoid trapping bubbles in the active cell volume. 14 µl is enough to fill the sample compartment and to avoid bubbles at the top of the window. Using more increases the chances of washing dust from the top part of the cuvette into the active volume of the cuvette.
17. Close the cuvette to prevent dust from getting in.
18. Wipe the cuvette from the outside with lens paper to remove any dust. Do not use Kimwipes it can scratch the quartz windows of the cuvette.
19. Insert the cuvette into the sample holder with the frosted side to the left!
20. Give the sample at least 5 minutes to equilibrate.
21. After equilibration check the readings that come in by clicking the Instrument Control Panel (icon to the right of the green light).
22. A new window opens displaying the incoming readings:
You can use this window to control laser power and acquisition time. Slowly increase laser power to 40%. If the values are higher than 2,000,000 cts/s disconnect immediately, these readings are high enough that they can damage the detector (avalanche photodiode)! A clean cuvette filled with filtered, bubble free H2O should have readings between 15,000 and 20,000 cts/s.
23. Repeat steps 15-22 with your filtered buffer.
24. If the values look consistent start data acquisition by clicking the green icon . It will turn red and blinking as the data acquisition is starting.
25. Do at least 10 acquisitions for the buffer control. If the values look good you can stop by clicking the now red icon.
26. If the data is good you can rinse the cuvette with filtered water and absolute EtOH and dry it as before. You are now ready to measure your sample.
27. If you plan to use more than one cuvette, test them all.
28. If the data looks suspicious and you are sure your buffer is dust and contamination free, clean the cuvette as described and try again. If you suspect your buffer is the issue, test with filtered Milli-Q water.
Sampling
29. For best results use the same cuvette for buffer and samples that you want to directly compare.
30. Load your sample in a dried, clean cuvette using a gel loading tip, avoiding bubbles as described above.
31. Close the cuvette, insert it into the sample holder with the frosted side to the left and give it 5 minutes to equilibrate.
32. Check the readings that come in by looking at the Instrument Control Panel.
33. A new window opens displaying the incoming readings. Remember, You can use this window to control laser power and acquisition time.
34. If changing the laser power does not give you a good range for the readings, you have to decrease (to high values) or increase (to low values) your sample concentration.
35. If the readings look good, start data acquisition. Do at least 20 acquisitions. Sometimes tiny bubbles can diffuse through the light path and skew some of the acquisitions. You can exclude those from the data evaluation later.
36. If you experience difficulties with your sample but the water control was fine, try re-filtering or re-centrifugation. Make sure your buffers are degased to avoid bubbles.
37. Sometimes it helps to simply tap the cuvette gently a couple of times on the bench to dislodge tiny bubbles.
38. Before you shut everything off, do at least 10 acquisitions of the cuvette after cleaning. Fill the cuvette with filtered H2O to obtain 10 data acquisitions proving you left the cuvette clean and save it as “Water_After_Cuvette #”.
39. Below is a typical result measured with 2mg/ml BSA, 40% laser power at 20 ℃.
40. The red marked readings were excluded from evaluation. The high values indicate an air bubble. Right click outlier acquisition. Select Mark. Selected acquisition will turn red, signifying exclusion.
41. View the Scattering Intensities
42. View Correlation Function
43. View Size Distribution
11.Western Blot
Introduction
Western blotting is used to visualize proteins that have been separated by gel electrophoresis. The gel is placed next to a nitrocellulose or PVDF (polyvinylidene fluoride) membrane and an electrical current causes the proteins to migrate from the gel to the membrane. The membrane can then be probed by antibodies specific for the target of interest, and visualized using secondary antibodies and detection reagents.
Procedure
Sample lysis
Preparation of lysate from cell culture
1. Place the cell culture dish on ice and wash the cells with ice-cold PBS.
2. Aspirate the PBS, then add ice-cold lysis buffer (1 mL per 107 cells/100 mm dish/150 cm2 flask; 0.5 mL per 5x106 cells/60 mm dish/75 cm2 flask).
3. Scrape adherent cells off the dish using a cold plastic cell scraper, then gently transfer the cell suspension into a pre-cooled microcentrifuge tube. Alternatively cells can be trypsinized and washed with PBS prior to resuspension in lysis buffer in a microcentrifuge tube.
4. Maintain constant agitation for 30 min at 4°C.
5. Centrifuge in a microcentrifuge at 4°C. You may have to vary the centrifugation force and time depending on the cell type; a guideline is 20 min at 12,000 rpm but this must be determined for your experiment (leukocytes need very light centrifugation).
6. Gently remove the tubes from the centrifuge and place on ice, aspirate the supernatant and place in a fresh tube kept on ice, and discard the pellet.
Preparation of lysate from tissues
1. Dissect the tissue of interest with clean tools, on ice preferably, and as quickly as possible to prevent degradation by proteases.
2. Place the tissue in round-bottom microcentrifuge tubes or Eppendorf tubes and immerse in liquid nitrogen to snap freeze. Store samples at -80°C for later use or keep on ice for immediate homogenization. For a ~5 mg piece of tissue, add ~300 μL of ice cold lysis buffer rapidly to the tube, homogenize with an electric homogenizer, rinse the blade twice with another 2 x 200 μL lysis buffer, then maintain constant agitation for 2 h at 4°C (eg place on an orbital shaker in the fridge). Volumes of lysis buffer must be determined in relation to the amount of tissue present; protein extract should not be too dilute to avoid loss of protein and large volumes of samples to be loaded onto gels. The minimum concentration is 0.1 mg/mL, optimal concentration is 1–5 mg/mL.
3. Centrifuge for 20 min at 12,000 rpm at 4°C in a microcentrifuge. Gently remove the tubes from the centrifuge and place on ice, aspirate the supernatant and place in a fresh tube kept on ice; discard the pellet.
Sample preparation
1. Remove a small volume of lysate to perform a protein quantification assay. Determine the protein concentration for each cell lysate.
2. Determine how much protein to load and add an equal volume 2X Laemmli sample buffer.
We recommend reducing and denaturing the samples using the following method unless the online antibody datasheet indicates that non-reducing and non-denaturing conditions should be used.
3. To reduce and denature your samples, boil each cell lysate in sample buffer at 100°C for 5 min. Lysates can be aliquoted and stored at -20°C for future use.
Loading and running the gel
1. Load equal amounts of protein into the wells of the SDS-PAGE gel, along with molecular weight markers. Load 20–30 μg of total protein from cell lysate or tissue homogenate, or 10–100 ng of purified protein.
2. Run the gel for 1–2 h at 100 V.
The time and voltage may require optimization. We recommend following the manufacturer’s instructions. A reducing gel should be used unless non-reducing conditions are recommended on the antibody datasheet.
Transferring the protein from the gel to the membrane
The membrane can be either nitrocellulose or PVDF. Activate PVDF with methanol for 1 min and rinse with transfer buffer before preparing the stack. The time and voltage of transfer may require some optimization. We recommend following the manufacturer’s instructions. Transfer of proteins to the membrane can be checked using Ponceau S staining before the blocking step.
Prepare the stack as follows:
Antibody staining
1. Block the membrane for 1 h at room temperature or overnight at 4°C using blocking buffer.
2. Incubate the membrane with appropriate dilutions of primary antibody in blocking buffer. We recommend overnight incubation at 4°C; other conditions can be optimized.
3. Wash the membrane in three washes of TBST, 5 min each.
4. Incubate the membrane with the recommended dilution of conjugated secondary antibody in blocking buffer at room temperature for 1 h.
5. Wash the membrane in three washes of TBST, 5 min each.
6. For signal development, follow the kit manufacturer’s recommendations. Remove excess reagent and cover the membrane in transparent plastic wrap.
7. Acquire image using darkroom development techniques for chemiluminescence, or normal image scanning methods for colorimetric detection.
12.BCA Assay
Prepare diluted albumin (BSA) standards:
1.Dilute the BSA standard into several clean vials, preferably using the same diluent as the samples.
2.Use the following table as a guide to prepare a set of BSA standards. Each 1 mL of BSA standard is sufficient to prepare a set of diluted standards for either working range suggested in the table.
Prepare BCA working reagent (WR):
1. Use the following formula to determine the total volume of WR required for the assay:
(standards + unknowns) × (replicates) × (volume of WR per sample) = total volume WR
Example: For the standard test-tube protocol with 3 unknowns and 2 replicates of each sample: (9 standards + 3 unknowns) × (2 replicates) × (2 mL) = 48 mL WR required
Note: For the test tube protocol, 2.0 mL of WR is required for each sample, while only 200 µL of WR reagent is required for each sample in the microplate procedure.
2. Prepare WR by mixing 50 parts of BCA reagent A with 1 part of BCA reagent B (50:1, Reagent A:B).
Note: When reagent B is first added to reagent A, turbidity is observed that quickly disappears on mixing to yield a clear, green WR.
Prepare sufficient volume of WR based on the number of samples to be assayed. The WR is stable for several days when stored in a closed container at room temperature.
Microplate procedure (sample to WR ratio = 1:8):
1. Pipette 25 µL of each standard or unknown sample replicates into a microplate well (working range = 20–2000 µg/mL). (For example, Thermo Scientific™ Pierce™ 96–Well Plates, Cat. No. 15041).
Note: If sample size is limited, 10 µL of each unknown sample and standard can be used (sample to WR ratio = 1:20), however, the working range is limited to 125–2,000 µg/mL.
2. Add 200 µL of the WR to each well and mix plate thoroughly on a plate shaker for 30 seconds.
3. Cover the plate and incubate at 37°C for 30 minutes.
4. Equilibrate the plate to room temperature. Measure the absorbance at or near 562 nm on a plate reader.
Note:
• Wavelengths from 540–590 nm have been used successfully with this method.
• Plate readers use a shorter light path length than cuvette spectrophotometers, hence the microplate procedure requires a greater sample to WR ratio to obtain the same sensitivity as the standard test tube procedure. If higher 562 nm measurements are desired, increase the incubation time to 2 hours.
• Increasing the incubation time or ratio of sample volume to WR increases the net 562 nm measurement for each well and lowers both the minimum detection level of the reagent and the working range of the assay. As long as all standards and unknowns are treated identically, such modifications are useful.
5. Subtract the average 562 nm absorbance measurement of the blank standard replicates from the 562 nm measurements of all other individual standard and unknown sample replicates.
6. Prepare a standard curve by plotting the average blank–corrected 562 nm measurement for each BSA standard vs. its concentration in µg/mL. Use the standard curve to determine the protein concentration of each unknown sample.
Note: If using curve-fitting algorithms associated with a microplate reader, a four-parameter (quadratic) or best–fit curve provides more accurate results than a purely linear fit. If plotting results by hand, a point–to–point curve is preferable to a linear fit to the standard points.
Apotome Fluorescence Microscope
1. Turn on the power switches in the following sequence: HXP120 (only if you need), Colibri control unit, ApoTome (only if you need), microscope power supply, microscope toggle, and computer. Shutdown sequence will be the opposite.
•Enter the Windows by logging on your username and password, then click the "Axiovision Rel. 4.8" icon to open the software.
•Load your specimen slide onto the stage, select the appropriate objective lens and focus the point of interest.
• Change lenses can be controlled by touch screen, Axiovision or manually.
• Don't switch from the oil lens to the dry lens. If it happens, remember to clean the dry lens!
• Make sure the transmitted light shutter is Open by checking the microscope, touch screen or Axiovision.
2. To obtain single bright field (BF), DIC or PH image (using transmitted light)
• Click "Workarea" to open the microscope/camera/processing window. Under the tab "Microscope", click the tab "Transmitted Light" and make sure the light is "On" with appropriate voltage.
•Select appropriate condenser turret manually or on touch screen. Focus the specimen, and adjust the slider if required, via eyepieces to get the best imaging effect.
•Click "Cam side port" and select the 100% light to the side.
•Click "Live", a window will pop up with the rough image of your specimen.
•Click "Property" to open a histogram, and adjust the quality of the image by playing with "Min/Max", "BestFit" and "Gamma" etc.
•Click "Snap" to capture the image.
•Click "Save" to save your image. Always save as ".zvi" format.
•To edit the images, click "Scale bar" etc. to label, measure or change colors etc.
3. To obtain single fluorescent image (using LED modules)
•Select the appropriate LED module and filter set based on the excitation/emission property of your dye/compound.
•Under the tab "Microscope", click the tab "Reflected Light".
•Under the "LED" list, click "desired LED On" or press the Colibri control panel wheel.
•Under the "Reflector" list to select the reflector (filter set).
•Make sure the FL (reflected light) shutter is "On" and TL "Off".
•Following above procedures to acquire and save images.
4. To obtain single fluorescence image (using HXP120)
•Turn on the light by switching on the power and pushing the Shutter button IN on the HXP120 unit. If this light source has just been turned off, please wait at least 20 minutes before you start it again!
•Click "Open" External Shutter under the "Colibri", or press "Ext"/"Shutter" on the Colibri control panel.
•Choose the appropriate filter under the "Reflector" list.
•Then following the same procedures as for LEDs to acquire and save images.
5. To obtain multiple images or multi-channel images (transmitted light and/or fluorescence) in the format of 2D, Z-stack or time-lapse
• Click the "6D Acquisition" button; or under "Workarea", click "Multidimensional Acquisition".
• Click the experiment we have saved previously.
•Click the "C" tab, set the channels you want to use.
•Then click "Measure" to optimize the exposure time for EACH individual channel. "OK" to accept the measurement.
•Click "Start" at the bottom to acquire multiple images.
•Following common steps to save and edit the images.
•You can set "Z-stack" or "Time Series" for each channel at your will by clicking the tab and following the on-screen instructions
6. To obtain single or multiple images with Apotome
•ApoTome power supply must be on when you start the microscope.
•First, following the same procedures as described above to acquire the ordinary single image.
•Then, carefully push the Apotome slider into the microscope.
•Click the "Apotome" icon on the left-hand menu. You can check if the grid (structured illumination in ApoTome) was calibrated with your lens and dye used.
•Click "Live" to acquire the image and snap/process as normal operations.
•The ApoTome slider should be off after use. To switch off ApoTome, just pull out the slider one step (hear a click-sound).
7. To obtain image with Extended Focus module
• Obtain normal Z-stack images under multi-dimensional mode and save. Select "Extended focus" under the tab "Processing" to open the saved file. It will convert the Z-stack images to a single sharp image.
• Or acquire the extended focus image by the "Extended Focus" module in the "Work area". Obtain the "live" image, manually adjust different focal planes and meantime click the "start" button. The images will automatically fuse to single extended focus format.
14.Apoptosis Assay by Flow Cytometry Using Annexin V Staining Method
This assay is used to count the number of cells that have undergone apoptosis. Apoptosis will be detected by initially staining the cells with Annexin V and propidium Iodide solution followed by flow cytometry analysis. It is based on the principle that normal cells are hydrophobic in nature as they express phosphatidyl serine in the inner membrane (side facing the cytoplasm) and when the cells undergo apoptosis, the inner membrane flips to become the outer membrane, thus exposing phosphatidyl serine. The exposed phosphatidyl serine is detected by Annexin V, and propidium iodide stains the necrotic cells, which have leaky DNA content that help to differentiate the apoptotic and necrotic cells.
Materials and Reagents
1.Annexin V FLUOS staining kit ( F. Hoffmann-La Roche, catalog number: 11858777001 )
2.The kit contains ready-to-use Annexin-V-FLUOS solution, propidium iodide solution, and incubation buffer
3.Trypsin
4.NaCl
5.KCl
6.Na2HPO4
7.KH2PO4
8.PBS buffer (pH 7.4)
Equipment
1.Flow cytometer
2.Centrifuge
3.T25 culture flask
Procedure
1.Seed cells (1 × 106 cells) in a T25 culture flask (in triplicate for experiments) and three T25 culture flasks for control (unstained, Annexin only, and propidium iodide only).
2.After 48 h incubation, collect the supernatant (floating apoptotic cells) and trypsinize the adherent cells (~2 × 106 cells) from each T25 flask (combine respective floating and trypsinized cells resulting in six tubes).
3.Wash the collected cells twice with PBS and centrifuge (670 × g, 5 min, RT).
4.Re-suspend each pellet (~2 × 106 cells) in PBS (400 µl).
For experimental cells (Triplicate):
(400 µl of cells + 100 µl of incubation buffer with 2 µl of Annexin [1 mg/ml] and 2µl of propidium iodide [1 mg/ml]).
For control cells:
Control 1: (unstained) - (without any stain (400 µl of cells + 100 µl of incubation buffer)
Control 2: (Annexin V only) - (400 µl of cells + 100 µl of incubation buffer with 2 µl of Annexin (1 mg/ml))
Control 3: (propidium iodide only) - (400 µl of cells + 100 µl of incubation buffer with 2 µl of propidium iodide (1 mg/ml))
5.Analyze the cells using a flow cytometry without washing the cells.
Cells that were propidium iodide (PI) negative and Annexin V negative are considered healthy, cells, PI negative and Annexin V positive cells are considered apoptotic, and cells that are positive to both PI and Annexin V considered necrotic.
15.Cell Proliferation Assay by IncuCyte (Sartorius)
Day 0
1. Plate Cells
1.1 Seed your choice of cells (100 μL per well) at an appropriate density into a 96-well plate, such that by day 1 the cell confluence is approximately 10–20%. The seeding density will need to be optimized for the cell line used; however, we have found that 1,000 to 2,500 cells per well (10,000 to 25,000 cells/mL seeding stock) are reasonable starting points.
• Monitor cell growth using the Incucyte® Live-Cell Analysis System to capture phase contrast images every two hours and analyze using the integrated confluence algorithm.
Day 1
2. Add Treatments (The isolated exosomes).
2.1 Prepare 1X concentrations of desired cell treatments in cell culture medium. The volumes may be varied; however, we recommend preparing enough volume of each desired treatment dilution in order to achieve 100 μL per well.
2.2 Remove the cell plate from the incubator and aspirate medium from wells.
2.3 Add treatments and controls to appropriate wells of the 96-well plate.
2.4 Place the cell plate into the Incucyte® Live-Cell Analysis System and allow the plate to warm to 37° C for 30 minutes prior to scanning.
a. Objective: 4X, 10X or 20X
b. Channel selection: Phase Contrast (+ Fluorescence if fluorescent label or cell health reagents are used)
c. Scan type: Standard
d. Scan interval: Typically, every 1 to 2 hours, until your experiment is complete. Note: Label free cell counting can be enabled on Incucyte® Live-Cell Analysis System with use of the Incucyte® Cell-by-Cell Analysis Software Module.
• Scan type: Standard | Adherent Cell-by-Cell
• Objective: 10X
Examples:
References
Enderle, D., Spiel, A., Coticchia, C. M., Berghoff, E., Mueller, R., Schlumpberger, M., Sprenger-Haussels, M., Shaffer, J. M., Lader, E., Skog, J., & Noerholm, M. (2015). Characterization of RNA from Exosomes and Other Extracellular Vesicles Isolated by a Novel Spin Column-Based Method. PLoS ONE, 10(8), e0136133.
https://doi.org/10.1371/journal.pone.0136133Erbse, A., Kethley, N., & University of Colorado. (n.d.). Basic Protocol for Dynamic Light Scattering.
https://www.colorado.edu/lab/biochem-instruments/sites/default/files/attached-files/dls_protocol_2017FACS analysis of fluorescent proteins - 2010.igem.org. (n.d.).
Incucyte®️ Proliferation Assays for Live-Cell Analysis | 연구용제품 > Bio마켓 | BRIC. (2023, August 9). BRIC.
https://www.ibric.org/bric/biomarket/product.do?mode=view&articleNo=9506097&title=IncucyteKessler, R. J., & Fanestil, D. D. (1986). Interference by lipids in the determination of protein using bicinchoninic acid. Analytical Biochemistry, 159(1), 138–142.
https://doi.org/10.1016/0003-2697(86)90318-0Lakshmanan, I., & Batra, S. K. (2013). Protocol for Apoptosis Assay by Flow Cytometry Using Annexin V Staining Method. Bio-protocol, 3(6), e374.
https://doi.org/10.21769/bioprotoc.374Shelke, G. V., Lässer, C., Gho, Y. S., & Lötvall, J. (2014). Importance of exosome depletion protocols to eliminate functional and RNA containing extracellular vesicles from fetal bovine serum. Journal of Extracellular Vesicles, 3(1).
https://doi.org/10.3402/jev.v3.24783Suchman, E. & American Society for Microbiology. (2016). Polymerase Chain Reaction Protocol.
https://asm.org/ASM/media/Protocol-Images/Polymerase-Chain-Reaction-Protocol.pdf?ext=Wiechelman, K. J., Braun, R. D., & Fitzpatrick, J. D. (1988). Investigation of the bicinchoninic acid protein assay: Identification of the groups responsible for color formation. Analytical Biochemistry, 175(1), 231–237.
https://doi.org/10.1016/0003-2697(88)90383-1
