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The following is a summary of our lab procedures.

Cell Culture

MCF-7, MDA-MB231, MDA-MB468, T47D, and HEK 293T cells were cultured in DMEM complete medium (10% FBS, 1% penicillin/streptomycin). HCC-1806 cells were cultured in RPMI-1640 complete medium (10% FBS, 1% penicillin/streptomycin), and MCF10A cells were cultured in commercially specialized medium.

The cells were maintained in a $\mathrm{CO}{}_2$ atmosphere at a constant temperature of 37°C, with humidity above 90%, in a $\mathrm{CO}{}_2$ incubator.

All cell experiment operations were performed in a sterile, toxin-free biosafety cabinet, ensuring the cells were kept clean, dust-free, and dry to avoid contamination.

All operations of cells were carried out in a sterile and non-toxic ultra-clean bench, and the cells were kept clean and dry to avoid contamination.

Cell Recovery

1. Disinfect the biosafety cabinet for 30 minutes, and turn on the thermostatic sterilized water bath. Preheat the medium to be used.
2. Take out the frozen cells from the -80°C ultra-low temperature freezer or liquid nitrogen, and immediately place them in a 37°C thermostatic sterilized water bath. Gently shake the cryovial to accelerate the thawing process, ensuring the thawing time does not exceed 1 minute.
3. Add 500 µL of complete medium to the cryovial, mix thoroughly by pipetting, and centrifuge at room temperature at 800 rpm for 5 minutes.
4. Discard the supernatant, gently resuspend the cells in 1 mL of complete medium, and transfer the cells to a 6 cm culture dish containing 4 mL of medium. Incubate at 37°C in a $\mathrm{CO}{}_2$ incubator.
5. After 12 hours, replace the medium, and perform cell passaging on the second day.

Cell Passage

1. Disinfect the biosafety cabinet for 30 minutes, and turn on the thermostatic sterilized water bath. Preheat the medium to be used.
2. When the cells in the culture dish reach 85% confluence, completely aspirate the medium and wash once with PBS.
3. Thoroughly aspirate the remaining PBS, add an appropriate amount of trypsin, and gently shake to cover all the cells. Digest in a 37°C $\mathrm{CO}{}_2$ incubator.
4. Under the microscope, when the edges of the digested cells appear transparent, the cells become round, and some cells detach, add complete medium to stop the digestion. Gently pipette the cells to detach most of them from the dish and transfer them to a centrifuge tube.
5. Centrifuge at room temperature at 800 rpm for 3 minutes.
6. Completely aspirate the supernatant, add an appropriate amount of complete medium, gently pipette to create a single-cell suspension, and transfer the cells to a culture dish with complete medium. Incubate at 37°C in a $\mathrm{CO}{}_2$ incubator. Replace the medium after 12 hours.

Cryopreservation of Cells

1. Disinfect the biosafety cabinet for 30 minutes, and turn on the thermostatic sterilized water bath. Preheat the medium to be used.
2. Repeat cell passaging steps (2) to (5).
3. Thoroughly aspirate the supernatant, add 1 mL of commercial serum-free cell freezing solution or laboratory-prepared cell freezing solution. Gently pipette to create a single-cell suspension. Transfer the cell suspension to cryovials and place them in a gradient cooling freezing box filled with isopropanol. Store overnight in a -80°C ultra-low temperature freezer, then transfer to a liquid nitrogen tank for long-term storage the next day.

Cell Counting

1. Disinfect the biosafety cabinet for 30 minutes, and turn on the thermostatic sterilized water bath. Preheat the medium to be used.
2. Repeat cell passaging steps (2) to (5).
3. Resuspend the digested cells in 1 mL of complete medium, gently pipette to mix and create a single-cell suspension. Take 50 µL of the cell suspension and add it to a 1.5 mL EP tube containing 950 µL of PBS, then gently pipette to mix.
4. Vortex to mix thoroughly, then use a small flow cytometer (Muse Cell Analyzer, Luminex Cooperation) to perform cell counting, with a dilution factor of 20x.

Protein Extraction

1. Use an ice box to pre-cool the NP-40 lysis buffer and PBS needed for the experiment, as well as the protease inhibitor at room temperature.
2. Aspirate the culture medium from the cultured cells and wash three times with pre-cooled PBS.
3. Prepare the lysis buffer containing proteinase inhibitors, and add it to the culture dish according to the cell quantity. Lyse the cells on ice for 15 minutes.
4. Use a cell scraper to detach the cells, transfer the cell lysate to a 1.5 mL EP tube, and sonicate at 30% power for 5 seconds, then pause for 5 seconds, repeating for a total of 5 cycles to achieve complete lysis.
5. Pre-cool the centrifuge to 4°C and centrifuge at 12,000 g for 15 minutes. Collect the supernatant and measure the protein concentration.

BCA Method for Determining Protein Concentration

1. Using the BCA protein quantification reagent, calculate the number of samples and the volume of working solution needed. Mix BCA Reagent A and Reagent B in a 50:1 ratio, which means 50 volumes of Reagent A to 1 volume of Reagent B. For example, mix 10 mL of Reagent A with 200 µL of Reagent B to prepare 10.2 mL of BCA working solution. The BCA working solution should be prepared fresh and not stored overnight.
2. Add standard samples of 0, 1, 2, 4, 8, 12, 16, and 20 µL into the wells of a 96-well plate to prepare a standard curve for protein. Add the corresponding protein lysate to each well to make the total volume 20 µL in each well. At this point, the protein concentrations in each well will be 0, 0.025, 0.05, 0.1, 0.2, 0.3, 0.4, and 0.5 mg/mL, respectively.
3. Add an appropriate amount of the protein sample to be tested into the wells of the 96-well plate, with each protein having three replicates. Add the corresponding protein lysate to make the total volume 20 µL in each well.
4. Add 200 µL of BCA working solution to the standard curve wells and the sample wells, and incubate at 37°C for 30 minutes (if the color is too light, the incubation can be extended to 60 minutes; conversely, if the color is too dark, reduce the incubation time to 20 minutes).
5. Measure the absorbance at 562 nm using an enzyme marker. Plot the standard curve of absorbance versus protein concentration, calculate the goodness of fit (R² value). An R² value greater than 0.99 indicates that the curve is acceptable. Compare the absorbance of the sample to the standard curve to determine the protein concentration of the sample.

Preparing SDS-PAGE Gel

1. Wash the glass plates (both long and short) with distilled water, then rinse with double-deionized water, and place them in a 55°C drying oven. Assemble the glass plates, secure them in the gel casting frame, and after confirming proper alignment, prepare the gel solution as follows:

Regant	8% Separation Gel	10% Separation Gel	12% Separation Gel	5% Concentrated Gel
30% acrylamide / bis-acrylamide solution (Acr/Bis) 29:1	2.7 mL	3.3 mL	4.0 mL	1.0 mL
Double deionized water	4.6 mL	4.0 mL	3.3 mL	4.1 mL
1.5 M Tris-HCl buffer (pH 8.8)	2.5 mL	2.5 mL	2.5 mL	/
1 M Tris-HCl buffer (pH 6.8)	/	/	/	0.75 mL
10% sodium dodecyl sulfate (SDS)	0.1 mL	0.1 mL	0.1 mL	0.06 mL
10% ammonium persulfate (APS)	0.1 mL	0.1 mL	0.1 mL	0.06 mL
Gel polymerization initiator	0.006 mL	0.004 mL	0.004 mL	0.006 mL

2. Calculate the total volume based on 8 mL of gel solution for each plate. Prepare the lower separating gel and pour the gel solution into the glass plates. Gently and evenly add anhydrous ethanol sealing gel, removing any bubbles generated during the pouring process while leveling the liquid surface.
3. After allowing the lower gel to set at room temperature for 30 minutes and observing a clear interface between the separating gel and ethanol, pour out the excess anhydrous ethanol and thoroughly absorb the remaining ethanol with absorbent paper.
4. Once the anhydrous ethanol is completely air-dried, add the upper concentrated gel. Simultaneously, slowly insert the comb along one side to avoid generating bubbles.
5. After allowing the upper gel to set at room temperature for 30 minutes, the gel should be fully solidified. The prepared gel can be stored in the electrophoresis buffer at 4°C or used immediately.

SDS-PAGE Gel Electrophoresis

1. Prepare the electrophoresis tank, electrodes, buffer, and gel. Assemble the gel and electrodes, then pour the buffer into the inner and outer tanks up to the marked line.
2. Slowly remove the comb and gently flush out any debris from the wells using a pipette tip.
3. Mix the protein sample with loading buffer, heat at 96°C for 10 minutes, and cool before loading the sample.
4. Run electrophoresis at 80V constant voltage for 30 minutes, ensuring the bromophenol blue bands fully enter the separating gel and the protein marker separates. Then switch to 120V constant voltage for 1 hour until the bromophenol blue bands reach the bottom of the gel.
5. After electrophoresis, proceed to either transfer or directly stain the gel with Coomassie Brilliant Blue or silver staining.

Semi-Dry/Wet Transfer Membrane Development

1. Prepare PVDF membrane, methanol, transfer buffer, ice box, and ice in advance.
2. Activate the PVDF membrane with methanol, then soak it in the transfer buffer.
3. Transfer conditions:
   • Wet transfer: constant voltage of 100 V for 90 minutes.
   • Semi-dry transfer: constant voltage of 25 V, limited to 1 A for 17 minutes.
4. After transfer, block the membrane with 5% BSA or 5% non-fat dry milk at room temperature for 1 hour.
5. Dilute the primary antibody and add it to the membrane after blocking. Incubate at room temperature for 2 hours or overnight at 4°C.
6. After primary antibody incubation, place the membrane on a shaker and wash it with TBS-T for 5 minutes, repeating the wash 4 times.

Coomassie Brilliant Blue Staining

1. Prepare Coomassie Brilliant Blue staining solution and destaining solution. Place the SDS-PAGE gel on a shaker and stain for 20 minutes.
2. After staining, recover the staining solution and destain using the Coomassie Brilliant Blue destaining solution. Change the destaining solution every hour until the bands are clear and the background is clean.
3. After destaining, take a photograph of the gel for analysis.

Agarose Gel Electrophoresis

1. Prepare agarose gel: Clean the agarose gel casting frame and a comb of appropriate dimensions. Assemble the casting frame.
2. Take a clean 250 mL triangular glass bottle, weigh 1 g of agarose powder, pour it into the bottle, and add 1 × TAE buffer to a final volume of 100 mL, preparing a 1% agarose gel (the gel concentration can be adjusted between 0.8% and 1.2% based on the size of the DNA molecules).
3. Place the triangular bottle in a microwave and heat on high for 2 minutes. After removing it, add Goldview nucleic acid dye, gently shake the bottle to ensure even mixing, and pour the solution into the assembled casting frame.
4. Allow the agarose gel to sit in the dark at room temperature for 30 minutes until it solidifies. Remove the comb and prepare for electrophoresis.
5. Agarose gel electrophoresis: Prepare 2 L of 1 × TAE buffer, pour it into the horizontal electrophoresis tank, and place the gel into the tank. Gently tap the gel to expel any air bubbles from the sample wells. Mix the DNA samples with the loading buffer evenly, and slowly add it to the sample wells, avoiding turbulence that could cause the samples to overflow. After loading, add a DNA marker to the wells on both sides of the samples.
6. Electrophorese at a constant voltage of 120 V for 30 minutes, or at 150 V for 20 minutes. Once electrophoresis is complete, place the gel into an ultraviolet imaging system for photography and analysis.

Plasmid Construction

1. Disinfect the biosafety cabinet for 30 minutes and prepare the target gene for amplification: Use commercially available target gene fragments or cDNA fragments amplified by reverse transcription, and prepare the PCR reaction system in a PCR tube according to the following formula:

Target Gene Fragment	1 µg
Forward Primer (10 µM)	1 µL
Reverse Primer (10 µM)	1 µL
2 × Pfu PCR Mix	25 µL
Double Deionized Water	Fill until 50 µL

2. After preparation, conduct regular PCR amplification according to the following procedure: (Repeat (1) to (3) for 35 times)

95°C Pre-denaturation	10 minutes
(1) 95°C Denaturation	30 seconds
(2) 58°C Annealing	30 seconds
(3) 72°C Extension	1000 bp/minute
72 °C Final Extension	5 minutes
4°C Refrigerated Storage	Indefinite

3. If constructing a point mutation plasmid, in addition to the forward and reverse primers, design the point mutation primers FM and RM:

F-end Fragment System:

Target Gene Fragment	1 µg
Forward Primer (10 µM)	1 µL
Reverse Primer (10 µM)	1 µL
2 × Pfu PCR Mix	25 µL
Double Deionized Water	Fill until 50 µL

R-end Fragment System:

Target Gene Fragment	1 µg
Forward Primer (10 µM)	1 µL
Reverse Primer (10 µM)	1 µL
2 × Pfu PCR Mix	25 µL
Double Deionized Water	Fill until 50 µL

Follow the regular PCR program for fragment amplification.

After amplification, perform a bridging reaction using the F and R fragments, as described below: (Repeat (1) to (3) for 8 times)

After completing the bridging reaction, add 1 µL each of the forward primer (10 µM) and the reverse primer (10 µM) to the system, and conduct another round of PCR. The product will be the amplified target gene fragment, which can be directly subjected to agarose gel electrophoresis for recovery, or temporarily stored at -40 °C.

DNA Purification / Gel Recovery

1. Sterilize the ultraclean workstation for 30 minutes, then equilibrate the spin column: Add 500 µL of Buffer BL to the spin column, let it stand for 1 minute, then centrifuge at 12,000 rpm for 1 minute. Discard the waste liquid in the collection tube and place the spin column back into the collection tube. The spin column should be processed the same day.
2. Gel recovery:
   1. Run agarose gel electrophoresis at a constant voltage of 150 V for 20 minutes. After electrophoresis, take the gel under UV light and cut out the target band (work quickly to avoid prolonged UV exposure that can break DNA in the gel). When cutting the gel, cut along the edge of the band to minimize leftover white gel (remove parts of the gel that do not contain DNA as much as possible). Place the cut gel piece into a pre-sterilized 1.5 mL EP tube, and weigh the cut gel piece. The weight of gel in each tube should not exceed 700 mg.
   2. Add 1 µL of Buffer MB per 1 mg of gel to the gel piece, mix, and place it in a 55°C metal bath at 300 rpm, shaking for 10 minutes. Every 3 minutes, invert the gel piece to speed up dissolution (when dissolving DNA fragments of 5 kb or longer, handle gently to avoid breaking long-chain DNA). Once the gel is dissolved, remove it from the metal bath.
3. DNA fragment binding: After the gel solution cools to room temperature, transfer the liquid into the equilibrated spin column with collection tube, let it sit at room temperature for 1 minute, and then centrifuge at 12,000 rpm for 1 minute. Discard the waste liquid in the collection tube and place the spin column back into the collection tube (make sure not to centrifuge in a low-temperature environment, as it can reduce binding efficiency). Repeat the previous step with the liquid in the collection tube (this step can improve recovery efficiency). If the volume of the solution exceeds 900 µL, which is more than the maximum capacity of the spin column, transfer the gel solution in two portions to the same spin column.
4. Column washing: Add 600 µL of Buffer MW containing 100% ethanol to the spin column, centrifuge at 12,000 rpm for 30 seconds, discard the waste liquid in the collection tube, and place the spin column back into the collection tube.
5. Repeat step (4) once, then centrifuge at 12,000 rpm for 1 minute to air-dry for 4 minutes to completely remove residual ethanol.
6. Place the spin column in a new 1.5 mL EP tube, add 35 µL of pre-heated (55°C) Endo-Free Buffer EB to the center of the membrane, let it sit at room temperature for 5 minutes, then centrifuge at 13,000 rpm for 2 minutes. Transfer the liquid from the EP tube to the spin column, repeat the process, and after centrifuging again at 13,000 rpm for 2 minutes, measure the particle concentration using a NanoDrop spectrophotometer for further experiments, or store at -40°C.

Digested Vector and Target Gene Fragment

Use appropriate restriction endonucleases to digest the amplified target gene fragments and suitable vectors from the above steps. Prepare the digestion reaction mixture in a PCR tube as follows:

Place the prepared digestion system in a 37°C water bath for digestion. After 3 hours of digestion (some endonucleases may require overnight digestion), perform agarose gel electrophoresis to recover the digested vector and target gene fragments.

Connection of Vector and Target Gene Fragment

After digestion, the connection of the vector and target gene fragments is usually performed using T4 ligase. Incubate at room temperature for 2 hours or at 16°C overnight. For sticky ends, the molar ratio of vector to target gene fragments is typically 1:7. This molar ratio yields a higher ligation efficiency. Prepare the ligation working solution according to the following ligation system:

Plasmid Transformation

1. Disinfect the ultra-clean bench for 30 minutes, prepare ice in a foam box.
2. Take the competent cells from the -80°C ultra-low temperature freezer and thaw them on ice for 5 minutes.
3. For newly ligated plasmids, take 15 μL of the ligation product and add it to 35 μL of competent cells. Gently mix by pipetting up and down, and incubate on ice for 30 minutes. For mature plasmids, take 100 ng and add it to 15 μL of competent cells, mix gently by pipetting, and incubate on ice for 30 minutes.
4. Preheat a water bath to 42°C. Heat shock the competent cells from the previous step in the water bath for 90 seconds, then immediately transfer them back to ice and incubate for 2 minutes.
5. Add 500 μL of antibiotic-free LB medium to the heat-shocked competent cells. For newly synthesized plasmids, place the mixture in a bacterial shaker at 220 rpm and 37°C for 1 hour for recovery.
6. Based on the resistance of the transformed plasmid, select the appropriate LB agar plates. For newly synthesized plasmids, plate 250 μL of the culture per plate. For mature plasmids, plate 100 μL of the culture per plate.
7. Incubate the plates at 37°C in an incubator for 14 hours and observe the formation of bacterial colonies.

Protein Purification

1. Construct the protein expression plasmid for purification.
2. Transform the plasmid into BL21(DE3) competent cells, following the plasmid transformation steps in Plasmid Transformation.
3. Select a single colony, grow the bacteria at 37°C for 8 hours, save 500 μL of the culture, collect 500 μL for sampling, and induce 1 mL of culture with 0.5 mM IPTG (or optimize using a gradient concentration) at 18°C for 20 hours.
4. Run the collected bacterial protein on SDS-PAGE, stain with Coomassie Brilliant Blue, following the staining steps in Coomassie Brilliant Blue Staining.
5. Observe whether the protein is successfully induced. For successful colonies, grow them overnight at 37°C in two 50 mL centrifuge tubes to expand the culture to 80 mL. The next day, add the expanded culture into 1 L of medium and grow at 37°C for 6 hours.
6. Induce with IPTG once the culture cools to 18°C. Add IPTG at a final concentration of 0.5 mM (or the optimal concentration determined in previous steps) and induce for 20 hours.
7. Centrifuge at 12,000 rpm for 2 minutes to collect the cells. Resuspend the collected bacterial culture in Protein Purification Buffer A (final volume less than 50 mL), place it in an ice-water mixture, add protease inhibitors (no more than 25 mL at a time), and use an ultrasonic disruptor to lyse the cells (parameters: 5 seconds on, 3 seconds off, power at 55%, for 99 cycles).
8. Centrifuge at 18,000 rpm at 4°C for 20 minutes, pour the supernatant into a new centrifuge tube.
9. Repeat the previous centrifugation step once more.
10. Filter the supernatant obtained using a 0.22 μM filter. Use an AKTA avant system and GST column for protein purification, then wash and elute with Protein Purification Buffer B.
11. Concentrate the eluted protein using an Amicon® Ultra filter (10 kDa MWCO), centrifuge at 4,000 rpm at 4°C until the desired volume is reached. Measure the protein concentration using a NanoDrop spectrophotometer, mix with 30% final concentration of glycerol, and store at -80°C in an ultra-low temperature freezer.

Medicine Screening

We completed high-throughput drug screening in our lab, conducted by a technician and a PRCXI SC9210 fully automated liquid handling workstation. The compounds used for screening were obtained from the MCE compound library, specifically the MCE novel known active compound library (1220 compounds). All compounds from the library were dissolved in 100% DMSO or H2O. By pre-diluting the drugs, a stock solution with a concentration of 1 mM was prepared.

The cells used for screening were MCF-10A normal breast epithelial cells and the MDA-MB-231 triple-negative breast cancer cell line. Cells were seeded in 96-well culture plates, with a cell concentration of about 5000 per well and a total volume of 100µL. After 24 hours of adherence, 1 μL of each compound from the library was aspirated and added to the culture medium of the 96-well plate using a fully automated four-dimensional modular processing platform, resulting in a final drug concentration of 10 µM. The negative control in this experiment was a pure DMSO group at 10 µM, while the positive control was a cisplatin group at 10 µM. After 48 hours of drug treatment, the technician discarded the culture medium in the wells, added ATP detection reagents, and performed luminance detection using a microplate reader, or ATP probe effect detection using a Revvity high-content imaging system.