Team:Jiangsu High School/Experiments
Experiments
Plasmid Transformation (Jul. 18)
Background:
In molecular biology, a vector is an artificial medium that carries foreign DNA material from one cell to another in order for it to be expressed or replicated in the latter. There are four major types of vectors: plasmids, viral vectors, cosmids, and artificial chromosomes; of which plasmids are the most common.
A couple of characteristics define a vector, such as having an origin of replication (where the vector could replicate itself autonomously inside its host), possessing a compatible site for insertion of genetic material, having the ability to clone in bulk, and retaining genetic markers that would predict if it is inside of its host (and use this to screen for non-recombinant cells).
Plasmids, the vector we used in this experiment, are generally circular, dual-stranded DNA molecules and are independent from the chromosomal DNA of any cell (because it possesses a piece of genetic material also known as the origin of replication). Plasmids could be in the cytoplasm of the cell but could not survive outside of its host. Plasmids can be transferred from one cell to another by bacterial conjugation. Not only could plasmids replicate itself independently, it could also integrate itself into its host's genomic information and replicate its material through the cell (known as transformation, where a cell adopts new genetic material from a vector).
When cells are placed in a 0 degrees Celsius CaCl2 solution, E. coli bacteria would become more permeable to the plasmid DNA. Then, after the cells are placed on ice along with the DNA, the solution would get heat-shocked at 42 degrees Celsius for about 2 minutes. This would promote the absorbance of the genetic material. The plasmid would then give the bacteria resistance towards a specific type of antibiotics, which could be used in the screening process.
Aim:
The aim of this experiment is to transform normal E. coli into competent cells with TagRPF-mWasabi-LC3 (the plasmid DNA).
Materials
• 1 working station
• 2 Erlenmeyer flasks
• Tap water
• Deionized Water
• 6 pieces of weighing paper
• 1 beaker
• Tin foil
• 1 Autoclave sterilizer
• 10g of tryptone
• 5g of yeast extract powder
• 10g of NaCl
• 50mg of ampicillin
• 10 large petri dishes
• 5 microliters of plasmid DNA
• 1 1mL pipette
• 1 5mL pipette
• 1 box of 1mL pipette tips
• 1 box of 5mL pipette tips
• 1 warm water bath
• 1 ice maker
• 1 ice box
• 1 centrifuge
• 1 waste jar
• 1 glass spreading rod
• 1 constant temperature incubator
• 8 1.5mL centrifugal tubes
Methodology:
• Thoroughly wash two erlenmeyer flasks first with tap water, then cleanse the walls with deionized water.
• Fold the weighing paper multiple times in order to maximize the surface area.
• Put the paper onto the balance and zero it.
• Make the mixture for the LB liquid culture medium by measuring out 10g of tryptone, 5g of yeast extract powder, and 10g of NaCl. Make sure to change out the weighing paper in between the different substances.
• Next, make the mixture for the LB solid culture medium by using the liquid culture medium, and 15g of agar powder. If the experimenters made a 2x solution, then they would use 500mL of H2O to dilute the solution.
• When both solutions are done, fill up a new beaker with 2L of deionized H2O. Distribute the water into the two flasks.
• Add tin foil to the openings of both flasks and place them into the autoclave sterilizer. Lock the device and set it to liquid mode. Press start. Let the apparatus run for 20 minutes.
• Add 50mg of ampicillin to the LB solid culture medium after the sterilizer. The reason we chose ampicillin is because the bacteria we are using (E. coli) are resistant whereas other bacteria that may exist within the culture are not.
• Deliver the LB solid culture medium using into 10 separate large petri dishes once the culture medium has cooled to 55 degrees Celsius.
• Add 5 microliters of plasmid DNA into 100 microliters of competent cells. Mix by pipetting the solution. This creates an environment with both DNA and cells.
• Place the new solution on ice for 30 minutes.
• After, place the tubes onto a float in a water bath that is 42 degrees Celsius for 42 seconds. This would allow the cell walls to become more porous and take in the plasmid DNA.
• Take the tubes out of the bath and back on ice for 2 to 3 minutes. The cold temperature would came the pores within the cell walls to constrict, thus trapping the DNA within each cell.
• For each tube of the new solution, add 1mL of LB liquid culture medium with no ampicillin. This is because after heating and cooling, adding a medium with ampicillin would damage the cells.
• Now, place the tubes into a shaking incubator for 45 minutes at 150rpm. Set the temperature to 37 degrees Celsius.
• Insert the tubes into the centrifuge and set the latter to 5000rpm for 2 minutes.
• Use a 1mL pipette to draw 700 microliters from the top layer of the tubes. Deliver the contents into the waste jar.
• Mix the 300 microliters with the cells on the bottom of the tube using a pipette.
• Using a 5mL pipette, deliver the solution in the tube into the petri dishes with the solidified culture medium. Use a glass spreading rod to make sure the solution covers the entire base of the dish and no bubbles are present.
• Let the dish sit for 15 minutes to allow the solution to be absorbed into the medium before inverting the dishes and placing them into a constant temperature incubator at 37 degrees Celsius overnight.
• The next day, the culture should be visible to the human eye. The bacteria colonies that are separate from the others are the E. coli with the plasmid DNA.
• Get a 50mL centrifugal tube and pour 30mL of liquid culture medium into said tube.
• Use a pipette to deliver 30 microliters of A+ solution into the tube. Place the pipette tip into the waste basket.
• Use an iron rod to select a lone colony of bacteria in the petri dish. Carefully dig out the colony and place it into the solution.
• Cap the tube and using a marker, write the experimenter's name and the plasmid name (LC3).
• Place the tube in the shaking incubator until the solution is well combined.
Cell Incubation (Jul. 18)
Aim
The aim of the experiment is to cultivate SH-SY5Y bacteria cultures for later research and experimentation. Even though SH-SY5Y cannot completely imitate body conditions as neural cells could, the experimenters chose the former because the latter is not possible to cultivate at an efficient pace, not possible to choose between specimens, and not suitable for later experiments involving autophagy.
Materials
• 1 tube of SH-SY5Y
• 1 liquid nitrogen container
• 1 water bath
• DMEM/F12
• DMSO
• FBS
• PS
• 1 5mL pipette
• 1 box of 5mL pipette tips
• 1 centrifuge
• 8 15mL centrifugal tubes
• 8 medium-sized petri dishes
• 1 marker
• 1 incubation box
Methodology
• Take out a tube of SH-SY5Y from a liquid nitrogen container. Then, preheat the water bath to 37 degrees Celsius.
• Swirl the tube in the water bath until the contents are fully liquefies.
• Make a solution of culture medium where 79% of DMEM/F12 (which contains glucose that would aid culture growth), 10% of DMSO (toxic to cells in large quantities under room temperature but normally used to prevent ice crystal growth during the freezing of the cells), 10% of FBS (fetal bovine serum, a blood fraction that remains after coagulation of blood [also didn't remove any remaining red blood cells [we use this in order to imitate the conditions within a living body and to stimulate culture growth]) and 1% of PS.
• Use a pipette to deliver the SH-SY5Y into a 15mL centrifugal tube and change to a 5mL pipette to deliver 4mL of the culture medium solution.
• Place the tubes in the centrifuge while making sure the apparatus is balanced. Set the centrifuge to 1000rpm for 2 minutes.
• After taking out of the tubes from the centrifuge, pour out the solution in the tube to leave the white sediments (the cells) at the bottom.
• Using a different pipette tip, deliver 1mL of culture medium solution into the centrifuge tube. Pipette the solution until it is thoroughly combined.
• Deliver all of the contents of the tube to a medium petri dish.
• Using the 5mL pipette, deliver 4mL of culture medium solution into the petri dish.
• Lightly swirl the petri dish in order to mix the solution evenly.
• Use a marker to write SH-SY5Y, the name of the experimenter, the date, and the generation number (F1 in this case).
• Place the petri dish into the incubation box.
细胞株与细胞系的区别
细胞株 (cell strain)
• A subpopulation of a cell line selected from the culture after undergoing cloning or some other method
• Euploid (a subculture that has an exact number of the haploid number of chromosomes)
• Incapable of indefinite serial passage
细胞系 (cell line)
• First subculture of a cell population of a primary culture
• Aneuploid (to have extra copies of the haploid number of chromosomes)
• Capable of indefinite growth
Plasmid Amplification, Extraction, and Purification (Jul. 20, 2019)
Background
This experiment would use alkaline lysis in order to extract plasmid DNA. In the pH range of 12-12.5, the double-helix structured DNA would undergo denaturation. In contrast, the hydrogen bonds within the circular-shaped plasmid DNA would break, but the two complementary strands would still be connected. When added pH 4.8 acid potassium acetate to lower the pH of the solution, the plasmid DNA would undergo renaturation whereas genomic DNA would not because of the bigger size of its molecules. Using the centrifuge, most of the cell debris, genomic DNA/RNA, and protein would become sediments at the bottom of the tube. All of the plasmid DNA would be in the supernatant. Using the cation column, isopropanol to induce sediments, and ethanol to rinse the solution, the experimenters would extract the purified plasmid DNA.
Aim
The aim of this experiment is to amplify the plasmid DNA via the shaking incubator, extract the plasmid DNA from the other proteins that might be present in the solution, and purify the solution in order to isolate the plasmid DNA.
Materials
• Centrifuge
• Waste jar
• 1 20 microliters pipette
• 1 box of 20 microliters pipette tip
• 1 500 microliters pipette
• 1 box of 500 microliters pipette tip
• 1 200 microliters pipette
• 1 box of 200 microliters pipette tip
• 150 microliters of P1 solution
• 150 microliters of P2 solution
• 350 microliters of P5 solution
• 1 absorption column
• 1 collector tube
• 300 microliters of PWT rinsing liquid
• 20 microliters of TB eluting buffer
• 1 EP tube
• 1 nucleic acid protein detector
• 2 microliters of buffer
• 1 marker
• 1 -20 degrees Celsius fridge
Methodology
• Before using PWT rinsing liquid, add absolute ethyl alcohol into the solution. Record volume added onto the bottle label.
• Take the bacterial solution out of the shaking incubator. Using a centrifuge, set it to 5000 rpm for 15 minutes.
• When done, deliver the supernatant into the waste jar.
• Add some deionized water into the tube. Use the pipette to thoroughly mix the solution.
• Place the tube into the centrifuge at 12000 rpm for 1 minute.
• Add 150 microliters of P1 solution which would wipe out the DNA polymerases. Make sure that RNA polymerase and TIANRed (a visual indicator, make sure the ratio of TIANRed to P1 solution is 1:200) are added into P1 before using.
• Use a pipette to thoroughly mix the solution until the sediments at the bottom of the tube is suspended in the solution. If the solution is not thoroughly mixed, this would affect quantity of solution that could be extracted later on and lower the purity of the solution.
• Add 150 microliters of P2 solution into the tube; the solution would damage the membranes and nuclei of the cells. Slowly shake the tube 6-8 times in order for the cells to be dissociated completely. At this time, the bacterial solution should be viscous and clear. If the solution is not clear, it might be because there are too much bacteria in the solution and the dissociation is not complete. If so, reduce the biomass of the bacteria in solution. After the P2 solution is thoroughly mixed in the tube, the solution should be a clear purple.
• Add 350 microliters of P5 solution into the tube in order to induce the plasmid DNA to undergo renaturation. Here, the genomic DNA would be completely damaged, leaving only the plasmid DNA. Quickly shake the solution until no purple color remains in the solution. Make sure all of the solution is yellow instead of purple to ensure the renaturation is successful. There will be cotton-like sediments in the tube.
• Place the tube into the centrifuge at 12000 rpm for 2 minutes.
• Use a pipette to deliver the supernatant (which now contains the plasmid DNA) into an absorption column with a collector tube underneath. Make sure not to pipette the sediments in the centrifugal tube.
• Place the absorption column into the centrifuge at 12000 rpm for 30 seconds.
• Pour away the solution in the collector tube and reassemble the absorption column on top of said tube.
• Add 300 microliters of PWT rinsing liquid into the absorption column.
• Centrifuge the column for 1.5 minutes at 12000 rpm in order to eliminate the remnants of PWT inside the absorption column.
• Pour away the solution in the collector tube into the waste jar and reassemble the absorption column.
• Use a pipette to deliver 20 microliters of TB eluting buffer into the absorption tube.
• Place the tube into the centrifuge at 12000 rpm for 30 seconds in order to collect the plasmid DNA in the collector tube.
• Use a pipette to deliver the plasmids DNA into a new EP tube.
• Measure the concentration of nucleic acid inside of the EP tube by first pipetting 2 microliters of buffer to zero the nucleic acid protein detector. Select DNA-50 and then Auto. Press Start.
• Pipette 2 microliters of the solution onto the instrument and then press Measure. The concentration should appear in units of ng/mL.
• Using a marker, write down the experimenter's name, the plasmid studied (LC3), the date, and the concentration of nucleic acid in solution.
• Place the labeled EP tube into a -20 degrees Celsius fridge for later use.
Developing Continuous Cell Lines (Jul. 20)
This method should be repeated every two days.
Aim
The aim of this practice is to give the cells more resources (both nutrients and space) to encourage more replication.
Materials
• 1 water bath
• 1 petri dish
• 1 5mL pipette
• 1 box of 5mL pipette tips
• 1 1mL pipette
• 1 box of 1mL pipette tips
• 1 waste jar
• 4mL of PBS
• 1mL of 0.25% pancreatic enzyme solution
• 1 thermal incubator
• 10mL of liquid culture medium
• 1 large petri dish
• 1 marker
Methodology
• Get the liquid culture medium, PBS (a buffer solution that would maintain the osmolarity of the cells [to balance the amount of salt ions in cells to prevent them from shrinking]), and pancreatic enzyme solution out of the fridge.
• Place all three solutions into a 37 degrees Celsius water bath.
• Get the medium-sized petri dishes from the incubator.
• Use a 5mL pipette to deliver the old liquid culture medium into the waste jar.
• Change the pipette tip.
• Use the 5mL pipette to deliver 4mL of PBS along the petri dish wall. Make sure to swirl the dish gently to allow the PBS buffer to rinse the cells.
• When done, deliver the used PBS buffer into the waste jar.
• Put the pipette tip into the waste jar.
• Use the 1mL pipette to deliver 1mL of 0.25% pancreatic enzyme solution which would reduce the ability of the cells to adhere to the dish walls. This would make it easier for the experimenters to collect as much cells for the transfer as possible.
• Gently swirl the solution around in the dish, making sure to cover not only the dish walls but also the base.
• When done, deliver the enzyme solution into the waste jar.
• Put the petri dish back into the incubator for 2 minutes for the cells to completely come off of the dish walls.
• Use the 5mL pipette to deliver 2mL of liquid culture medium into the petri dish. Use the pipette to collect the solution and pipette the solution over the entirety of the base to ensure all of the cells are submerged in the culture medium itself.
• Now, pipette the solution with the same tip into a large-sized petri dish. This would give more space for the cells to grow.
• Change the pipette tip.
• Because the volume of a large petri dish is 10mL, use the 5mL pipette to pipette 8mL of liquid culture medium into the dish.
• Using a marker, record down the experimenter, the generation number, the cell experimented with, and the date of the experiment.
• Now gently swirl the dish until there are no air bubble present in the solution.
• Place the dish in the incubator.
Cell Medium Change (Jul. 20)
This procedure should be repeated every other day.
Aim
The aim of this procedure is to ensure the cells have enough nutrients for them to continue their replication process.
Materials
• 1 box of 5mL pipette tips
• 1 5mL pipette
• 4mL of PBS rinsing solution
• 5mL of liquid culture medium
• 1 waste jar
Methodology
• Using a 5mL pipette, deliver the old liquid culture medium into the waste jar.
• Change the pipette tip.
• Add 4mL of PBS rinsing solution to the cell walls. Deposit into the waste jar using the pipette after swirling to rinse the cells.
• Change the pipette tip.
• Add 5mL of new liquid culture medium to allow cells to continue their replication process.
• Return the petri dish into the incubator.
Plasmid Transfection (Jul. 22)
Background
Transfection is a method of implanting foreign genetic material into a cell. There are different methods of transfection: for this experiment, we are using lipofection. Lipofection is association of nucleic acid with a cationic lipid solution. The resulting molecular complexes (called lipoplexes) then enter into the cells' genomic information. By entering the nucleus, transfection could then replicate its genetic material within its host.
Aim
The aim of this experiment is to use lipofection to implant LC3 into the cells.
Materials
• 1 cotton swab
• 1 bottle of 75% ethyl alcohol
• 1 5mL pipette
• 1 box of 5mL pipette tips
• 1 1mL pipette
• 1 box of 1mL pipette tips
• 4mL of PBS solution
• 1mL of 0.25% pancreatic enzyme
• 4mL of normal liquid culture medium
• 1 waste jar
• 1 centrifuge
• 1 15mL centrifugal tube
• 1 12-well cell culture cluster
• 2 EP tubes
• 150 microliters of Opti-MEM
• 2 microliters of Lip2000
• 500ng of plasmid DNA
• 800 microliters liquid culture medium with no PS nor FBS
Methodology
• Use 75% ethyl alcohol in order to disinfect your work station as well as your working gloves. Use a cotton swab to clean the working surface. When done, deposit the swab into the waste jar.
• Using a 5mL pipette, deliver the liquid culture medium into the waste jar.
• Change the pipette tip.
• Using the same pipette, deliver 4mL of PBS solution along the dish walls in order to rinse the petri dish. Gently swirl the dish around to make sure it covers the dish surfaces.
• Add 1mL of 0.25% pancreatic enzyme into the petri dish. Gently swirl the solution to cover all the dish surfaces.
• Place the petri dish back into the incubator for 1 minute to wait for the cells to dissociate.
• Add 2mL of liquid culture medium. Use the 1mL pipette to thoroughly mix the solution as well as pipette solution to make sure all of the cells are submerged in solution.
• Make sure to place glass slides inside the wells before injecting the cells inside the cells to make sure the cells could be taken out for the autophagy flux later.
• Deposit the pipette tip into the waste jar.
• Using the 5mL pipette, deliver the solution into a 15mL centrifugal tube.
• Set the centrifuge to 1000 rpm for 3 minutes.
• Add 1mL of the solution to 1mL of liquid culture medium. Mix thoroughly with pipette.
• Using a 1mL pipette, deliver 15 microliters of the solution from the tube per well into a 12-well cell culture cluster.
• After the cells adhered to the dish walls, add liquid culture medium that does not include PS nor FBS (only DMEM/F12).
• In a new EP tube, mix 75 microliters of Opti-MEM (a lipofection reagant) with 2 microliters of Lip2000. Remember to change the pipette tip in between different solutions.
• In another new EP tube, mix 75 microliters of Opti-MEM with 500ng of plasmid DNA.
• Carefully pipette the second tube's contents into the first. Wait 15 minutes.
• Add the new solution to the cell solution in the cell culture cluster.
• Wait for 6 hours to let the cells absorb the solution.
• After 6 hours, experimenters could use a fluorescence microscope to see if any of the cells absorbed the LC3 plasmid DNA.
• Using the 1mL pipette, add 1mL of regular liquid culture medium (89% DMEM/F12, 10% FBS, and 1% PS) to the wells.
• After 24 hours, the experimenters would add autophagy inhibitors (CQ and 3-MA) as well as three compounds (Compound 1, 2, and 3).
Preparation for Detection of Autophagy Flux (July 24)
Background
Macroautophagy is a process in which autophagosomes sought to degrade and "recycle" damaged proteins or impaired organelles after fusing with lysosomes to turn into autolysomes. Autophagy is being pushed further into the spotlight in the recent years because how its dysfunction may lead to many neurodegenerative diseases such as Alzheimer's Disease (AD) (Zhang et all, 2013). Damage in the autophagy mechanism is directly linked to the protein accumulation which would increase toxicity in neuro-pathways of animal models. Mutations in presenilin-1 caused primarily the failure of autophagy to breakdown protein; this would eventually lead to neuronal cell death in AD models (Lee et all, 2010).
Aim
The aim of today's experiment is to prepare the solutions to see the effectiveness of each well in response to the autophagy flux test which would happen the next day.
Materials
• 1 bottle of Hyclone SH30023.01
• 1 bottle of FBS
• 1 37 degrees Celsius water bath
• Compounds 1-8, OD 1-3, and OD 5-15
• 12 EP tubes
• 1 test tube holder
• 1 1mL pipette
• 1 box of 1mL pipette tip
• 1 5mL pipette
• 1 box of 5mL pipette tip
• 2 cell clusters
• 1 waste jar
• 1 incubator box
• 1 marker
Methodology
• Take out the Hyclone SH30023.01 DMEM/F12 and FBS from the fridge.
• Place them into the warm water bath at 37 degrees Celsius until the solutions thaw.
• From the fridge, take out Compound 1-8 as well as OD1-3 and 5-15.
• Clean your station with a cotton swab after dipping it in alcohol.
• Take out a 15mL centrifugal tube.
• Measure out the appropriate amount of 89% DMEM/F12, 10% FBS, and 1% PS. Be sure to change out the pipette tip in between delivering different solutions.
• Take out 12 EP tubes and set them onto the test tube holder.
• After mixing the liquid culture medium, use a 1mL pipette to deliver 1050 microliters of liquid culture medium solution to each EP tube.
• Do not put anything except for the liquid culture medium solution into the first EP tube. For the second one, add 1.05 microliters of DMEM in order to emulate the DMEM added to the compound solutions while making them.
• Add the rest of the compound into their respective tubes after DMEM has been added.
• Get the 12-well cell cluster from the incubator box. This cluster should have the solutions with the Compound 1-8 and OD 1-3. [3-MA: 自噬抑制剂。一直自噬小体的发生—no autophagosome=no autolysosome=没有红、绿/或者红和绿会被降弱]
• Using the 1mL pipette, deliver the supernatant into the waste jar.
• Change the pipette tip in between delivering different solutions.
• Transfer the contents of each tube into their respective wells after mixing thoroughly with your pipette.
• Place the lid of the cell cluster onto said instrument.
• Use a marker to specify the solution added to each well.
• Place the cluster into the incubator.
• Repeat the process with the second 12-well cell cluster that would include the solutions OD5-15.
Cell Preparation for Frozen Storage (July 24)
Aim
If cell samples would not be used for a period of time, they will be stored into long term storage (-80 degrees Celsius) to save space for samples that are more frequently used in the -20 degrees Celsius storage. This procedure is to prepare cell samples for long term frozen storage.
Materials
• 9.45mL of liquid culture medium
• 1.05mL of DMSO
• 1 medium-sized petri dish
• 1 incubator box
• Cell samples to be frozen
• 1 box of 5mL pipette tips
• 1 box of 1mL pipette tips
• 1 5mL pipette
• 1 1mL pipette
• 12mL of PBS
• 1 waste jar
• 6mL of 0.25% pancreatic enzyme
• 9 frozen storage tubes
• 1 marker
Methodology
• Prepare the frozen storage solution (90% liquid culture medium and 10% DMSO). In our case, we are making 3.5mL of storage solution so we will add 3.15mL of liquid culture medium and 0.35mL of DMSO. It is essential to add DMSO into the solution before it lowers the freezing point of the cells as well as prevents the cells from losing moisture and crystalizing (this would destroy the cell structure).
• Mix the frozen storage solution into a medium-sized petri dish.
• Take out the large petri dish from the incubator box.
• Using a 5mL pipette, deliver the supernatant into the waste jar.
• Change the pipette tip.
• Rinse the cells with 4mL of PBS. Make sure to add the PBS solution into the walls of the incubator.
• Slowly swirl the dish in order to cover all the surfaces.
• Deliver the PBS into the waste jar.
• Change the pipette tip.
• Deliver 2mL of 0.25% pancreatic enzyme to the dish in order to loosen the cells' grip on the walls of the dish.
• Deliver the pancreatic enzyme into the waste jar.
• Change the pipette tip.
• Place the dish into the incubator box for 2 seconds to speed up the process of the cells coming off of the dish walls.
• Deliver the frozen storage solution into the large petri dish.
• Tilt the dish and pipette solution from the far end of the dish to the near one, making sure the cells are coming down with the solution.
• After making sure most, if not all, of the cells are immersed in solution, take 3 frozen storage tubes out.
• Changing the pipette tip in between, deliver 1mL of solution into each tube.
• Using a marker, write SH-SY5Y, your name, and the date of the procedure.
• Above is the procedure for one petri dish. Repeat until all of the petri dishes are done.
Detection of Autophagy Flux (July 26)
Aim
The aim for today's experiment is to analyze the photos we got from the confocal microscope.
Materials
• 1 incubator
• Prepared cell clusters
• 1 5mL pipette
• 1 box of 5mL pipette tips
• 1 waste jar
• 1 bottle of PBS solution
• 1 bottle of 4% PFA solution
• 1 timer
• 1 bottle of Solarbio Hoechst 33342 stain
• 6 glass slides
• 1 container to place the glass slides in
• 1 marker
• 1 tweezer
• 1 confocal microscope
• 1 Leica computer app
Methodology
• This procedure is best done in a dark room with no direct light. Because the fluorescence inside the cells may be damaged from white light.
• From the incubator, take out the 12-well cell clusters.
• Using a 5mL pipette, deliver the liquid culture medium from the wells and into the waste jar.
• Change the pipette tip.
• Deliver PBS solution into each well in order to rinse the wells of damaged or dead cell material. Make sure the solution completely submerge the base of the well. Make sure the pipette tip is placed against the well's wall when delivering the solution.
• Swirl the solution gently.
• Using the same pipette tip, deliver the solution into the waste jar.
• Repeat this procedure again.
• Now, add 4% PFA solution in order to fix the cells onto the circular glass slides placed in the wells before the plasmid transfection. Make sure the pipette tip is held against the walls when delivering the solution. The solution should submerged the base of the wells.
• Wait for 10 minutes.
• Using a pipette, deliver the PFA solution into the waste jar.
• Change the pipette tip.
• Using a pipette, deliver PBS solution to the wells while making sure it covers the base. Gently swirl it and deliver the solution into the waste jar.
• Change the pipette tip.
• Add Solarbio Hoechst 33342 stain to stain the DNA just enough to cover the base of the wells as well.
• Wait for 10 minutes for the stain to settle.
• When the time is done, deliver the solution into the waste jar.
• Repeat the rinsing process of PBS stated above for 3 times to clean out the stain.
• In a microscope slide container, place down 6 glass slides (since 2 solutions could be placed per slide).
• Using a pipette, add a drop of anti-fade mounting medium for each circular slide to be placed on (2 drops per slide). Make sure to give space between the two drops since both has to be covered with circular slides.
• Use a marker to write down the solution the circular slides were placed on. Remember that the first well is the control group that did not have any compound added into its liquid cell culture medium.
• Use a tweezer to take out the circular slides from each well. After taking out the slide, be sure to rotate the slide 180 degrees so that the side with the cells would face downwards and be in contact with the anti-fade mounting medium.
• Continue this procedure until all of the circular slides are taken out and place on the glass slides in the container.
• Take the microscope slide container to the confocal microscope.
• Take out a slide. Make sure to rinse the lens with 75% ethanol with tissue paper before placing the slide on for examination.
• Use the knobs to adjust what you can see until you see the clearest and most (in number) cells. The more cells and the better they are stained, the more likely their other fluorescent tags would show.
• Using the Leica computer app, press "Live" at the bottom left corner to get the view from the microscope and you could use the knobs as well to adjust your field of vision. We have 3 different sequences (the Hoechst blue stain, the mWasabi green fluorescent light, and the red fluorescent light). For each one, pan around the field of vision until you've decided you have the best representation of the compound's authophagy mechanisms.
• Use the knob to adjust the lens distance to the dimmest point where you could still see the subject of your field of vision (for example, a stained nucleus). Click "Begin" on the left column and then begin to scroll until the other dimmest point, then click "end".
• Click "Start" at the bottom of the screen for the lens to start taking photos of the different layers of the cells on the slide. This may take a while.
• When the photographs are done, click the next tab on the top of the screen and then click "Project Tools". Click "Projection" and then "Apply".
• Click on the right column to select the different stains you want to adjust the brightness of. The objective of this is to get rid of background stains and to emphasize the essential reactions of the compound to autophagy. Click on "C1", "C2", and "C3" to separately adjust the vibrancy and brightness of the stains.
• When done, rename the project to your compound name to not confuse it with the other compounds.
• Repeat this process until you are done with all of the compounds in the microscopic slide box.