Polymerase Chain Reaction (PCR) is an effective laboratory technique for DNA amplification. It used to rapidly create millions to billions of copies of a specific segment of DNA.

1. Assemble all reaction components on ice. Each component should be gently mixed before adding to the reaction in a sterile thin-walled PCR tube. The entire reaction should be mixed again to ensure homogeneity. Collect all liquid to the bottom of the tube with a quick spin if necessary. Overlay the sample with mineral oil if using a PCR machine without a heated lid.
2. Add different components into the sterile thin-walled PCR tube according to the proportion below:

   1. First Round PCR

      10 μM Forward Primer	1μl
      10 μM Reverse Primer	1μl
      DNA	1μl
      One Taq 2X Master Mix with Standard Buffer	25μl
      Nuclease-free water	22μl

   2. Second Round PCR

      UTR mix	3μl
      First Round PCR Product	5μl
      One Taq 2X Master Mix with Standard Buffer	25μl
      Nuclease-free water	17μl

3. Put the tubes into the PCR machine and start it.

   Thermocycling condition for a routine PCR

   STEP	TEMP	TIME
   Initial Denaturation	95°C	30 seconds
   25-35 Cycles	95°C 45-68°C* 68°C	15-30 seconds 15-60 seconds 1 minute/kb
   Final Extension	68°C	5 minutes
   Hold	4-10°C

Transformation is the process of introducing foreign DNA — such as a plasmid into a host cell. We used BL21, a type of e-coli. to take up the plasmid and express new genetic material.

1. Thaw competent cells on ice.
2. Chill approximately 5 ng (2 μl) of the ligation mixture in a 1.5 ml microcentrifuge tube.
3. Add 50 μl of competent cells to the DNA. Mix gently by pipetting up and down or flicking the tube 4–5 times to mix the cells and DNA. Do not vortex.
4. Place the mixture on ice for 30 minutes. Do not mix.
5. Heat shock at 42°C for 30 seconds. Do not mix.
6. Add 950 μl of room temperature media to the tube.
7. Place tube at 37°C for 60 minutes. Shake vigorously (250 rpm) or rotate.
8. Warm selection plates to 37°C.
9. Spread 50–100 μl of the cells and ligation mixture onto the plates.
10. Incubate overnight at 37°C.

Gel electrophoresis is a commonly used laboratory technique that separates charged molecules—such as DNA, RNA, or proteins—based on their size. In this method, an electric field drives the molecules through a porous gel matrix. Smaller molecules migrate faster through the gel's pores, while larger ones move slower, resulting in distinct bands that can be visualized for analysis.

1. Preparation of agarose

   1. Measure 0.3 g agarose powder
   2. Mix agarose power with 30mL 1X TAE buffer with a microwavable flask
   3. Microwave solution for 1 minute. Microwave for longer if there is undissolved agarose powder
   4. Let the agarose solution cool down to about 50°C, about 5 minutes.
   5. Add 3µl GelGreen® Nucleic Acid Gel Stain into the gel solution, and mix them well.
2. Pour the solution into the gel tray. Insert the comb into the top of the gel.
3. Wait for around 15 minutes to let the gel solidify.

4. Running gel electrophoresis

   1. Once the gel has solidified, carefully remove the comb by pulling it straight up.
   2. Place the agarose gel into the gel box.
   3. Fill the gel box with 1 X TAE buffer until the gel is covered.
   4. Prepare the sample by adding a loading buffer to each, with the ratio of 1:5.
   5. Load 15μL of DNA ladder into the first lane of the gel.
   6. Load samples into wells. Avoid bubbles.
   7. Run the gel at 80-150 V until the dye line is approximately 75-80% of the way down the gel. A typical run time is about 1-1.5 hours, depending on the gel concentration and voltage.
   8. Turn off the power and carefully remove the gel from the gel box.
   9. Image it with UV.

Protein purification is a laborious process used to isolate a specific and targeted protein from a complex mixture.

1. Reagent preparation
   1. DTT Solution: Dissolve one vial of DTT (Solid) in 1 mL of ultrapure water. DTT solution must be prepared fresh before used
   2. Lysis Buffer: Mix Native Lysis Buffer with freshly prepared DTT solution at a 200:1 ratio.
   3. Wash Buffer: Mix Native Wash Buffer with freshly prepared DTT solution at a 200:1 ratio.
   4. Elution Buffer: Mix Elution Wash Buffer with freshly prepared DTT solution at a 200:1 ratio.
2. Add 20μL of Lysis Buffer into the bacteria with our DNA, and resuspend the bacteria in the lysate sufficiently, allowing for a slight vortex (to minimize bubbles).
3. Centrifuge at 4°C for 10 min.
4. Add 500 μL of 50% His tag Purification Resin (Reductant and Chelator Resin) mixed evenly, and slowly shake on a shaker at 4°C for 30 minutes to fully bind the target protein with His tag.
5. Centrifuge 10 s at 4°C to precipitate the gel, take 20 μL of the supernatant for subsequent detection, and discard the rest of the supernatant.
6. Add 100 μL Wash Buffer resuspension gel, centrifuge at 4°C 10 s, retention sample for subsequent detection, and discard the rest. Take 20 μL supernatant
7. Repeat step (6) with one more wash.
8. Add 20 μL of Elution Buffer and gently resuspend the gel. Centrifuge at 4°C 10 s to collect the supernatant and gel. The supernatant is the purified His-tagged protein of interest.
9. Repeat step (8) twice. Approximately 60 μL of purified protein samples were collected by co-elution.

SDS-PAGE is used to separate proteins based on their molecular weight. The proteins are loaded onto a polyacrylamide gel. When an electric field is applied, proteins migrate through the gel toward the positive electrode. We can identify the relative molecular weights by comparing the migration distance of proteins.

1. Place the gel in the SDS-PAGE stand
2. Place the stand with the gel in the SDS-PAGE apparatus bath
3. Fill the space between the gels with running buffer
4. Take off the combs of the gel (take it out gently)
5. Load the sample to each well
6. Do not use the wells at the left and right end of the gel
7. Fill the SDS-PAGE apparatus bath with running buffer to the bottom of the gel
8. Place the cover of the SDS-PAGE
9. Plug the cables to the SDS-PAGE apparatus power source
10. Turn off the SDS-PAGE power source
11. Set to constant voltage
12. Use 200 V
13. Press the Run button to start the electrophoresis
14. The voltage increases to 200 V from 0 V
15. For 4% staking gel and 12.5% separating gel the current reaches 50-60 mA and the power 10-12 W
16. The current then the power decreases in time but not reach zero (if it reaches zero see the troubleshoot)
17. For the state type of gel it takes 35 - 40 mins

1. Thaw competent cells on ice
2. Add 1-5μl (10pg-100 ng) of plasmid (do not exceed 5μl for a 50μl cell aliquot)
3. Incubate on ice for 30 minutes
4. Heat-shock by placing in 42°C water bath for exactly 30 seconds
5. Place cells on ice for 2 minutes
6. Add 1mL pre-warmed LB or SOC medium
7. Shake incubate 37°C, 200rpm, 1 hour for outgrowth

If monitoring cell density by OD600, record final OD readings prior to harvesting.

1. Harvest cells by centrifugation at 16,000 × g for 10 minutes. For larger volumes, centrifuge for ≥30 minutes especially if at ≤10,000 × g. Discard the medium and, if necessary, weigh the wet cell pellet.
2. Store the pellet at -20°C or -80°C or process immediately.
3. Resuspend the cell pellet in NEBExpress E. coli Lysis Reagent by pipetting or vortexing briefly until the suspension is homogenous:
4. Use 0.025 - 0.075 mL of NEBExpress E. coli Lysis Reagent for every 1 UOD600 harvested. To calculate the UOD600, multiply the volume harvested by the OD600 reading. For example, a 5 mL culture harvested at OD600 1 gives 5 mL x 1.0 = 5 UOD600. In this example, 0.125 – 0.375 mL Lysis Reagent is required to lyse efficiently.
5. If harvested cells are weighed, use 5 mL of NEBExpress E. coli Lysis Reagent per 1 gram of cells.
6. Incubate the resuspended cells at room temperature for 10 - 20 min with gentle shaking, gentle rotation, or swirling. Lysis is usually visible with a clearance of the suspension.
7. Centrifuge the lysate at 16,000 x g for 10 min at 4°C to pellet the insoluble material and cell debris (30 min or longer for large volumes and lower speed).
8. Carefully transfer the supernatant into a sterile container for analysis or purification. This soluble fraction can be stored at 4°C for a few hours or -20°C or -80°C for longer term storage.
9. If needed, resuspend the insoluble pellet in 50 mM

ELISA is a widely used laboratory method for detecting and quantifying specific substances—typically proteins, antigens, or antibodies—in various biological samples such as blood, serum, or cell culture supernatants.

Reagent Preparation

1. Dilute the wash solution with deionized water (1:20)
2. Set standard wells and test sample wells. Add 50 μl standard to standard well.
3. Add 10 μl testing sample then add 40 μl sample diluent to testing sample well; blank well does not add anything.
4. Add 100 μl of HRP conjugate reagent to each well, cover with an adhesive strip and incubate for 60 minutes at 37°C.
5. Aspirate each well and wash, repeating the process four times total of five washes, wash by filling each well with Wash solution (400 μl) using a squirt bottle, manifold dispenser or autowasher. Complete removal of liquid at each step is essential to good performance. After the last wash, remove any remaining Wash Solution by aspirating or decanting. Invert the plate and blot it against clean paper towels.
6. Add chromogen solution A 50μl and chromogen solution B 50μl to each well. Gently mix and incubate for 15 minutes at 37°C. Protect from light.
7. Add 50μl Stop solution to each well. The color in the wells should change from blue to yellow if the color in the wells is green or the color change does not appear uniform, gently tap the plate to ensure thorough mixing.
8. Read the Optical Density (O.D.) at 450nm using a microtiter plate reader within 15 minutes.

Calculation of Results

1. This standard curve is used to determine the amount in an unknown sample. The standard curve generated by plotting the average O.D. (450 nm) obtained for each of the six standard concentrations on the vertical (Y) axis versus the corresponding concentration on the horizontal (X) axis.
2. First, calculate the mean O.D value for each standard and sample. All O.D values, are subtracted by the mean value of the zero standard before result interpretation. Construct the standard curve using graph paper or statistical software.
3. To determine the amount each sample, first locate the O.D. value on the Y-axis and extend a horizontal line to the standard curve, t the point of intersection, draw a vertical line to the X-axis and read the corresponding concentration
4. Any variation in operator, pipetting and washing technique, incubation time or temperature, and kit age can cause variation in result. Each user should obtain their own standard curve.
5. The sensitivity by this assay is 1.0 pg/ml

Cell Revival

1. Preheat water bath to 37°C.
2. Prepare one 15 ml centrifuge tube, add 5 ml complete medium containing 10% FBS, and preheat in 37°C water bath.
3. Wear protective goggles and gloves, take out the frozen cells from liquid nitrogen quickly and place them in the preheated water bath to increase thawing speed.
4. Transfer thawed cells to the prepared centrifuge tube, mix well, and centrifuge at 1000 rpm for 5 minutes.
5. Prepare a T-25 flask, write cell name and date, add 4 ml complete medium.
6. After centrifugation, discard supernatant, resuspend the cell pellet in 1 ml complete medium, transfer to T-25 flask, and incubate in a CO₂ incubator.

*Caution:* When taking frozen cells from liquid nitrogen, if liquid nitrogen enters the cryovial, loosen the cap to release internal pressure, then quickly tighten and place on dry ice before placing in 37°C water bath to avoid explosion due to rapid nitrogen evaporation.

Cell Passage

1. Pass cells when confluency reaches over 85%.
2. In the biosafety cabinet, open the culture flask cap and collect the culture medium.
3. Add 3 ml sterile 1×PBS, lay flat to rinse cells, discard PBS.
4. Add 1 ml digestive solution to bottle, incubate at 37°C CO₂ incubator for 1-2 minutes.
5. Observe under inverted microscope if cells have detached; if fully digested, add 2 ml complete medium with 10% FBS, and aspirate into a 15 ml centrifuge tube.
6. (If cells are not fully digested, perform additional digestion steps in portions using specific steps described.)
7. Centrifuge at 1000 rpm for 5 min.
8. Prepare two new T-25 flasks, add 4 ml complete medium to each.
9. After centrifugation, discard supernatant, resuspend pellet in 2 ml complete medium, split evenly into two T-25 flasks (1 ml each).
10. Place flasks in a 37°C incubator with 5% CO₂ for static culture.

Cell Freezing

1. Follow steps 1-6 from the passage procedure.
2. After centrifugation and discarding supernatant, resuspend cells in 1 ml freezing medium to precipitate cells.
3. Transfer to 1.8 ml cryovials.
4. Place cryovials filled with DMSO in a programmable freezing container, then transfer to -80°C freezer overnight.
5. On the second day, transfer cryovials from freezing container into liquid nitrogen for long-term storage.

Testing NO production in trabecular meshwork cell lines:

I. Preparation of Working Solution

Take an appropriate amount of the stock solution and dilute it to the working concentration using the dilution buffer provided in this kit or self-prepared fresh culture medium (without serum and phenol red). For example, take 5 µL of DAF-FM DA (5 mM) and dilute it at a 1:1000 ratio to prepare 5 mL of DAF-FM DA (5 µM) working solution. Mix thoroughly.

[Note] The working solution must be prepared fresh for immediate use. Discard any excess probe after the experiment. Since the dye is more prone to oxidation and decomposition in aqueous solution, please use it as quickly as possible.

[Note] The optimal working concentration of the probe needs to be determined based on your specific cell type or experimental conditions, or by consulting relevant literature. Generally, choose the minimum probe concentration that ensures sufficient fluorescence signal, as this helps minimize the accumulation of ester hydrolysis byproducts such as formaldehyde and acetic acid.

II. Probe Loading

• For cells with short stimulation times (usually within 2 hours): load the probe first, then stimulate the cells with an appropriate positive control and your drug of interest.
• For cells with long stimulation times (usually more than 6 hours): stimulate the cells with an appropriate positive control and your drug of interest first, then load the probe.

1. In-situ Probe Loading (Applicable only to adherent cells)

   1. Remove the cell culture medium and add an appropriate volume of the prepared DAF-FM DA working solution (e.g., the working solution diluted 1:1000 times). The volume added should be sufficient to cover the cells adequately; typically, add 1 mL of working solution per well of a 6-well plate.
   2. Incubate at 37°C for 20 minutes.
   3. Wash the cells three times with PBS, pH 7.4, to thoroughly remove any DAF-FM DA that has not entered the cells.

2. Probe Loading After Cell Harvesting

   1. After harvesting the cells, resuspend them in an appropriate volume of the prepared DAF-FM DA working solution (e.g., the working solution diluted 1:1000 times) to achieve a cell concentration of 1-20 million cells per milliliter.
   2. Incubate at 37°C for 20 minutes. The above operations can be performed in a centrifuge tube. Invert and mix gently every 3-5 minutes to ensure full contact between the probe and cells.
   3. Wash the cells three times with PBS, pH 7.4, to thoroughly remove any DAF-FM DA that has not entered the cells.
   4. Directly stimulate the cells with an appropriate positive control or your drug of interest, or first divide the cells equally into several aliquots and then stimulate them.

III. Fluorescence Detection

1. For samples with in-situ loaded probe: You can observe directly using a confocal laser scanning microscope (observation with a standard fluorescence microscope yields relatively poorer results), or harvest the cells and detect using a fluorescence spectrophotometer, fluorescence microplate reader, or flow cytometer.
2. For samples where the probe was loaded after cell harvesting: You can detect using a fluorescence spectrophotometer, fluorescence microplate reader, or flow cytometer. Direct observation with a confocal laser scanning microscope is also possible.
3. Parameter settings: Use an excitation wavelength of 495 nm and an emission wavelength of 515 nm. Perform real-time, time-lapse, or single time-point detection of fluorescence intensity before and after stimulation. The fluorescence spectrum of the DAF-FM-NO reaction product is very similar to that of Fluorescein. You can use the parameter settings for detecting Fluorescein, or alternatively, use the parameter settings for detecting FITC.

IV. Additional Notes

1. The recommended working concentration for DAF-FM DA in the above steps is 5 µM. For some cell types, if the fluorescence of the unstimulated negative control cells is also relatively strong, you can dilute DAF-FM DA at a ratio of 1:2000 to 1:5000, so that the working concentration of DAF-FM DA during probe loading is 1-2.5 µM. Conversely, if the fluorescence is weak after stimulation with your drug of interest, you can adjust the working concentration of DAF-FM DA to 10 µM to improve detection sensitivity.
2. The probe loading time can be adjusted appropriately within the range of 15-60 minutes based on the specific situation.

Testing cell apoptosis rate in retinal ganglion cell lines

1. Induce cell apoptosis using the experimental protocol.

2. Collect the cells

   • Suspension cells: Centrifuge at 300 × g for 5 min and aspirate the culture medium supernatant.
   • Adherent cells: Trypsinize the cells, preferably using trypsin without EDTA. Centrifuge at 300 × g for 5 min and aspirate the supernatant.
3. Wash the cells with ice-cold PBS and collect 1-5 × 10⁵ cells.
4. Resuspend the cells in 500 µL of 1X Binding Buffer.
5. Add 5 µL of Annexin V-FITC and 5 µL of Propidium Iodide. Gently mix and incubate at room temperature for 10-20 minutes, protected from light.

6. Perform detection immediately after the reaction using a flow cytometer or fluorescence microscope. If immediate detection is not possible, keep the samples on ice and analyze within 1 hour.

   1. Flow Cytometry Detection:

      • Annexin V-FITC green fluorescence has an emission wavelength of 530 nm and can be detected using the FL1 (FITC detector) channel.
      • PI red fluorescence has an emission at 620 nm and can be detected using the FL2 or FL3 channel. Using the FL3 channel is recommended.

   2. Fluorescence Microscopy Observation:

      • Place a 30-50 µL drop of the cell suspension from Step 5 onto a glass slide and cover it with a coverslip. Observe under a fluorescence/confocal laser scanning microscope.
      • Cells positive for Annexin V-FITC staining (FITC filter, blue light excitation) will show bright green fluorescence on the cell membrane.
      • Cells positive for Propidium Iodide staining (PI filter, green light excitation) will show red fluorescence.
      • Early apoptotic cells are not stained by PI, while necrotic or late apoptotic cells show red cytoplasm, red nuclei, and a surrounding green cell membrane. Membrane shrinkage and blebbing can also be observed in late apoptotic cells.