General Protocols

Electroporation of Escherichia coli Nissle 1917

Introduction

Electroporation is a highly efficient method for introducing plasmid DNA into bacterial cells by applying a short, high-voltage pulse to temporarily permeabilize the cell membrane.

Procedures

1. Pick a single colony of E. coli Nissle 1917 from an LB agar plate and inoculate into 5 mL fresh LB broth. Incubate at 37 °C, 220 rpm for 12 h.
2. Dilute the overnight culture into three tubes, each containing 5 mL LB broth, at a 1:200 ratio. Grow at 37 °C, 220 rpm until OD₆₀₀ ≈ 0.3.
3. Place the cultures on ice for 10 min.
4. Centrifuge at 4 °C, 3000 × g, 10 min. Discard the supernatant and resuspend the cells in 3 mL ice-cold 10% glycerol.
5. Repeat Step 4 three times.
6. Centrifuge again at 4 °C, 3000 rpm, 10 min. Discard the supernatant and gently resuspend the pellet in 1 mL ice-cold 10% glycerol to obtain the competent cell suspension.
7. Aliquot the competent cells into five pre-chilled 1.5 mL microcentrifuge tubes, 110 µL per tube.
8. Add 10 µL plasmid DNA (approx. 1.5–2.0 µg) to 110 µL competent cells and mix gently.
9. Transfer the mixture into a pre-chilled 1 mm electroporation cuvette.
10. Electroporate at 2440 V with a pulse duration of 4–6 ms.
11. Immediately add 800 µL LB broth to resuspend the cells.
12. Transfer to a sterile tube, incubate statically at 37 °C for 30 min, then shake at 37 °C, 220 rpm for 2 h for recovery.
13. Centrifuge at 5000 × g, 2 min, discard the supernatant, and resuspend the cells in fresh LB broth.
14. Spread the suspension onto LB agar plates containing ampicillin.
15. Incubate at 37 °C for 12 h.

Preparation of Electrocompetent Cells and Electroporation of Staphylococcus epidermidis ATCC 14990

Introduction

Electroporation is a highly efficient method for introducing plasmid DNA into bacterial cells by applying a short, high-voltage pulse to temporarily permeabilize the cell membrane.

Procedures

A. Preparation of Electrocompetent Cells — Method 1

1. Inoculate a single colony of S. epidermidis into 25 mL B medium and grow at 37 °C with shaking (250 rpm) for ~24 h.
2. Dilute the overnight culture to OD₆₀₀ = 0.1 in fresh pre-warmed B medium. Grow at 37 °C, 250 rpm until OD₆₀₀ ≈ 2.0 (3–6 h).
3. Dilute the culture again to OD₆₀₀ = 0.1 and continue cultivation.
4. Harvest cells at early exponential phase (OD₆₀₀ = 0.5–0.65) by centrifugation at 5900 × g, 10 min, 4 °C.
5. Wash the pellet with ice-cold 10% glycerol, centrifuge (5900 × g, 10 min, 4 °C), and repeat three times with progressively reduced volumes of 10% glycerol (½, 1/20, 1/50 of the original culture volume).
6. Resuspend the final pellet in 700 µL ice-cold 10% glycerol. Aliquot 60 µL per tube. Store at −80 °C or use immediately.

A. Preparation of Electrocompetent Cells — Method 2

1. Inoculate a single colony of S. epidermidis into 10 mL BM (or TSB) medium and grow 8 h.
2. Transfer into 90 mL fresh pre-warmed BM/TSB and grow until OD₆₀₀ = 0.8–0.9.
3. Chill culture on ice for 10 min. Centrifuge at 3900 × g, 5 min, 4 °C.
4. Wash cells with 100 mL ice-cold sterile water, centrifuge at 3900 × g, 5 min, 4 °C.
5. Repeat previous step.
6. Finally resuspend it in 10% glycerol.
7. Aliquot 50 µL per tube. Store at −80 °C.

B. Electroporation of Plasmid DNA — Method 1

1. Prepare high-quality plasmid DNA (≥500 ng/µL, free of protein/RNA contamination).
2. Thaw competent cells (60 µL) at room temperature.
3. Add 300–1000 ng plasmid DNA, mix gently, incubate at room temperature for 30 min.
4. Transfer to a pre-chilled 1 mm electroporation cuvette. Electroporate at 2 kV, 25 µF, 100 Ω.
5. Immediately add 950 µL SMMP medium.
6. Recover at 37 °C, 250 rpm for 4 h (if using temperature-sensitive plasmids such as pE194 ts derivatives, incubate at 30 °C).
7. Centrifuge at 5000 × g, 2 min, discard the supernatant, and resuspend the cells in fresh LB broth.
8. Spread the suspension onto agar plates containing selected antibiotics.
9. Incubate at 37 °C for 12 h.

B. Electroporation of Plasmid DNA — Method 2

1. Thaw competent cells (50 µL) on ice for 5 min, then at room temperature for 5 min.
2. Centrifuge and resuspend in 10% glycerol + 500 mM sucrose.
3. Add 5 µg pre-purified plasmid DNA, transfer to a 1 mm cuvette (Bio-Rad). Electroporate at 21 kV/cm, 25 µF, 100 Ω.
4. Immediately recover cells in BHI medium supplemented with 500 mM sucrose at 28 °C for 2 h.
5. Centrifuge at 5000 × g, 2 min, discard most of the supernatant, and resuspend the cells by remaining medium.
6. Spread the suspension onto BHI agar plates containing chloramphenicol 10 μg/mL.
7. Incubate at 37 °C for 12 h.

Preparation of Electrocompetent Cells and Electroporation of Staphylococcus xylosus ATCC 29971

Introduction

Electroporation is a highly efficient method for introducing plasmid DNA into bacterial cells by applying a short, high-voltage pulse to temporarily permeabilize the cell membrane.

Procedures

1. Pick a single bacterial colony and inoculate it into 1 mL of BHI broth; incubate at 37 °C with shaking for 12 h.
2. Dilute the culture 1:100 into 50 mL of BHI broth and incubate at 37 °C with shaking until OD₆₀₀ reaches 0.4–0.5 (≈ 2.5–3 h).
3. Place the culture on ice for 10 min, then centrifuge at 2,800 × g, 4 °C, 10 min.
4. Resuspend the pellet in 30 mL ice-cold 10% glycine and incubate on ice for 5 min.
5. Sequentially wash the cells with 10 mL, 2.5 mL, and 1 mL ice-cold 10% glycine, centrifuging after each wash.
6. Resuspend the pellet in 200 μL ice-cold 10% glycine.

Note: Competent cells should be freshly prepared and used immediately.

7. Add 5 μg plasmid DNA (≤ 5 μL) to 200 μL competent cells; incubate at room temperature (25 °C) for 30 min.
8. Transfer 100 μL mixture into a pre-chilled 0.1 cm electroporation cuvette.
9. Apply the following pulse: 2 kV, 25 μF, 100 Ω; immediately add 1 mL pre-warmed BHI broth supplemented with 0.5 M sucrose (37 °C), mix gently.
10. Incubate at 37 °C with shaking for 1 h.
11. Centrifuge at 12,000 rpm (≈ 13,800 × g) for 1 min; discard the supernatant and retain ~100 μL of culture.
12. Spread the concentrated cells onto BHI + 0.5 M sucrose agar plates containing 12.5 μg/mL chloramphenicol.
13. Incubate at 37 °C for 24 h until single colonies appear.

Preparation of Electrocompetent Cells and Electroporation of Escherichia coli Nissle 1917

Introduction

Electroporation is a highly efficient method for introducing plasmid DNA into bacterial cells by applying a short, high-voltage pulse to temporarily permeabilize the cell membrane.

Procedures

A. Preparation of Electrocompetent Cells

1. Streak E. coli Nissle 1917 (EcN 1917) onto an LB agar plate and incubate at 37 °C for 12 h.
2. Inoculate a single colony into 5 mL LB medium and culture at 37 °C, 220 rpm for 12 h.
3. Dilute the overnight culture into 5 mL fresh LB medium at a 1:200 ratio. Grow until OD₆₀₀ = 0.3 (37 °C, 220 rpm).
4. Place the culture on ice for 10 min; centrifuge at 4 °C, 3000 rcf for 10 min to harvest the cells.
5. Wash the pellet three times with 3 mL ice-cold 10% glycerol, centrifuging after each wash.
6. After washing, resuspend the cells in 1 mL ice-cold 10% glycerol. Aliquot into 1.5 mL tubes (100 µL per tube) for immediate use in transformation. Alternatively, snap-freeze the aliquots in liquid nitrogen and store at –80 °C for later use.

B. Electroporation of EcN 1917 with Plasmid DNA

1. Add 10–15 µL plasmid DNA (≈1.5–2 µg) to 100 µL electrocompetent cells. Incubate the mixture on ice for 10 min.
2. Transfer the DNA-cell mixture into a pre-chilled electroporation cuvette (gap width 0.2 cm).
3. Electroporate at 3.0 kV (15 kV/cm), pulse time ~4 ms.
4. Immediately add 800 µL LB medium to the cuvette, resuspend cells, and transfer to a sterile tube.
5. Recover at 37 °C for 1–1.5 h with shaking.
6. Centrifuge at 6000 rcf for 1 min, discard ~600 µL of the supernatant, and resuspend the cells in the remaining medium.
7. Plate the suspension onto LB agar plates containing the appropriate selective marker. Incubate at 37 °C for 12 h.

Preparation of Chemically Competent Cells and Heat-Shock Transformation (CaCl₂ Method)

Introduction

The CaCl₂ method is a classical and widely used approach to prepare E. coli cells for transformation. By treating cells with cold calcium chloride, the cell membrane becomes more permeable to DNA molecules. This treatment neutralizes the negative charges on both the DNA and the cell surface, facilitating DNA uptake during heat-shock transformation. Although not as efficient as electroporation, the CaCl₂ method is simple, cost-effective, and sufficient for most routine cloning applications.

Procedures

A. Preparation of Chemically Competent Cells

1. Streak bacteria from a −80 °C stock onto LB agar plates. Incubate overnight at 37 °C.
2. Pick a single colony and inoculate 5 mL LB broth (with antibiotics if necessary). Incubate overnight at 37 °C with shaking at 200 rpm.
3. Transfer 500 µL of the overnight culture into 50 mL LB broth. Grow at 37 °C with shaking (200 rpm) until OD₆₀₀ ≈ 0.5. Immediately place culture on ice for 30 min.
4. Transfer culture into pre-chilled 50 mL centrifuge tubes. Centrifuge at 4 °C, 5000 rpm, 10 min. Discard supernatant.
5. Resuspend pellet in ~35–40 mL pre-chilled 0.1 M CaCl₂. Incubate on ice for 30 min.
6. Centrifuge at 4 °C, 4500 rpm, 15 min. Discard supernatant.
7. Resuspend pellet in 2–3 mL of ice-cold CaCl₂ solution containing 10% glycerol.
8. Aliquot into sterile 1.5 mL tubes (typically 100 µL per tube).
9. Snap-freeze aliquots in liquid nitrogen or dry ice/ethanol bath. Store at −80 °C until use.

B. Heat-shock Transformation of Competent Cells

1. Add the plasmid DNA or ligation product (not exceeding 10% of competent cell volume) into 100 µL competent cells. Mix gently and incubate on ice for 30 min.
2. Transfer the tube to a 42 °C water bath for 90 s. Immediately place the cells back on ice for 2 min.
3. Add 200 µL LB broth (no antibiotics) to the cells. Incubate at 37 °C, 200 rpm for 1 h to allow expression of antibiotic resistance.
4. Spread the culture onto LB agar plates containing the corresponding antibiotic. Incubate the plates overnight at 37 °C.

PCR Protocol: Thermo Scientific Phusion High-Fidelity PCR Master Mix

Introduction

The Polymerase Chain Reaction (PCR) is a widely used molecular biology technique that allows rapid and specific amplification of DNA fragments.

Procedures

Prepare a 20 µL or 50 µL reaction system as shown below, assembling all components on ice. Mix each reagent gently before adding it into a sterile thin-walled PCR tube. Ensure the entire reaction mixture is homogeneous; if necessary, briefly centrifuge to collect all liquid at the bottom of the tube.

Immediately transfer the reaction mixture to a thermal cycler preheated to the denaturation temperature and set the cycling parameters according to the following conditions.

PCR Protocol: 2X M5 HiPer plus Taq HiFi PCR mix (with blue dye)

Introduction

The Polymerase Chain Reaction (PCR) is a widely used molecular biology technique that allows rapid and specific amplification of DNA fragments.

Procedures

Prepare a 20 µL reaction system as shown below, assembling all components on ice. Mix each reagent gently before adding it into a sterile thin-walled PCR tube. Ensure the entire reaction mixture is homogeneous; if necessary, briefly centrifuge to collect all liquid at the bottom of the tube.

Immediately transfer the reaction mixture to a thermal cycler preheated to the denaturation temperature and set the cycling parameters according to the following conditions.

PCR Protocol: Phanta Flash Master Mix (Dye Plus)

Introduction

The Polymerase Chain Reaction (PCR) is a widely used molecular biology technique that allows rapid and specific amplification of DNA fragments.

Procedures

Prepare a 50 µL reaction system as shown below, assembling all components on ice. Mix each reagent gently before adding it into a sterile thin-walled PCR tube. Ensure the entire reaction mixture is homogeneous; if necessary, briefly centrifuge to collect all liquid at the bottom of the tube.

The optimal reaction concentration varies depending on the template. In a 50 µL reaction system, the recommended amount of template is as follows:

Immediately transfer the reaction mixture to a thermal cycler preheated to the denaturation temperature and set the cycling parameters according to the following conditions.

Protocol: Beyotime PCR Clean Up kit / DNA Purification Kit

Introduction

PCR purification or DNA purification kits are designed to efficiently remove primers, nucleotides, enzymes, salts, and other impurities from DNA samples. The principle is based on selective binding of DNA to a silica membrane in the presence of chaotropic salts, followed by washing to eliminate contaminants and elution of pure DNA in a low-salt buffer or water.

Procedures

1. Add an equal volume of Solution 1 to the sample and mix thoroughly.
2. Transfer the mixture (sample + equal volume of Solution 1) into a DNA spin column. Let it stand at room temperature for 1 min.
3. Centrifuge at maximum speed (16,000 × g, approx. 12,000–14,000 rpm) for 1 min. Discard the flow-through in the collection tube.
4. Add 700 µL Solution II into the spin column and let stand at room temperature for 1 min.
5. Centrifuge at maximum speed for 1 min to remove impurities. Discard the flow-through.
6. Add 500 µL Solution III, centrifuge at maximum speed for 1 min to further wash away impurities. Discard the flow-through.
7. Centrifuge again at maximum speed for 1 min to remove residual liquid and allow remaining ethanol to fully evaporate.
8. Place the spin column into a clean 1.5 mL microcentrifuge tube. Add 50 µL Elution Buffer (Solution I) directly to the membrane. Let stand for 1 min. (Optionally, use 20–30 µL for higher concentration but lower yield. Incubating for 3–5 min may slightly improve yield.)
9. Centrifuge at maximum speed for 1 min. The eluate contains the purified DNA.

Agarose Gel Electrophoresis

Introduction

Agarose gel electrophoresis is a widely used method for the separation and analysis of nucleic acids, such as DNA and RNA. The technique relies on an electric field to move negatively charged nucleic acid molecules through a porous agarose matrix, where fragments are separated according to size. Smaller fragments migrate more quickly, while larger fragments move more slowly. By comparing band patterns to a DNA ladder of known sizes, researchers can estimate fragment lengths, verify PCR or cloning results, and assess DNA purity.

Procedures

1. Measure 1.0 or 1.2 g of agarose. (Agarose gels are typically prepared at 0.7%–2% concentration depending on fragment size; for ~800 bp plasmids, 1%–1.2% works well.)
2. Mix agarose powder with 100 mL 1× TAE in a microwavable flask.
3. Microwave for 1–3 min until the agarose is completely dissolved. Avoid overheating to prevent evaporation that alters final gel percentage. Heat in 30 s intervals, swirl to mix, then continue until fully dissolved.
4. Let agarose solution cool down to about 50 °C (3–5 min).
5. Add 10 µL GelRed directly to the molten agarose before it solidifies and mix gently.
6. Pour the agarose into a gel tray with the well comb in place. Level the tray, pour slowly to avoid bubbles, remove bubbles with a pipette tip if needed.
7. Allow the gel to solidify at room temperature for 20–30 min (or place at 4 °C for faster solidification).
8. Once solidified, place the agarose gel into the gel box (electrophoresis unit).
9. Fill gel box with 1× TAE (or TBE) until the gel is fully covered.
10. Carefully load a molecular weight DNA ladder into the first lane (typical loading volume ~5 µL).
11. Carefully load your samples into the additional wells (typical loading volume ~10 µL).
12. Run the gel at 80–150 V until the dye front is ~75–80% of the way down the gel (usually 1–1.5 h depending on concentration and voltage). Lower voltage for better band resolution.
13. Turn off power, disconnect electrodes, and carefully remove the gel from the box.
14. Image the gel using a gel documentation system.

Protocol: Colony PCR for Screening of Transformants

Introduction

Colony PCR is a rapid and convenient method used to identify recombinant clones directly from bacterial colonies without the need for plasmid extraction. A small portion of each colony is used as the DNA template in a PCR reaction, allowing researchers to quickly verify the presence or absence of the desired insert or construct. This method greatly reduces screening time and is widely applied in molecular cloning to distinguish positive transformants from non-recombinant colonies.

Procedures

1. Prepare a 50 µL PCR reaction mixture on ice. Thoroughly mix each reagent before adding. Use sterile thin-walled PCR tubes and add components according to the manufacturer’s instructions. After assembly, gently mix the reaction mixture and, if necessary, briefly centrifuge to collect the liquid.
2. Select a well-isolated, healthy single colony from an LB agar plate. Mark its position on the plate and record the colony ID.
3. Using a sterile pipette tip or inoculating loop, first streak the colony onto a fresh antibiotic-containing plate (to establish a backup culture), then touch the same tip/loop to the PCR mixture as the DNA template source.
4. For Gram-positive bacteria, pick a single colony into 30 μL of NaOH (20 mM) solution and process it according to the following protocol: 95°C for 5 minutes; 4°C for 1 minute, repeat this cycle three times.
5. Gently stir a small amount of cells into the prepared PCR mixture (slight turbidity visible to the naked eye is sufficient), taking care not to carry over agar.
6. Place the PCR tube into a thermal cycler for amplification. Cycling parameters should be set according to the DNA polymerase used and the experimental purpose.

Protocol for Plasmid Miniprep Using Spin Column

Introduction

Plasmid miniprep is a standard method used to isolate and purify plasmid DNA from small volumes of bacterial culture. The spin column method uses alkaline lysis followed by silica membrane DNA binding and washing to obtain clean plasmid DNA suitable for downstream molecular biology applications.

Procedures

1. Add 1–5 mL of overnight bacterial culture to a 1.5 mL or 2 mL centrifuge tube. Centrifuge at 12,000 × g for 1 min and discard the supernatant.
2. Add 250 µL Buffer S1 to the cell pellet. Resuspend completely until no clumps remain.
3. Add 250 µL Buffer S2, gently invert the tube 4–6 times until the solution becomes clear and viscous. Do not vortex.
4. Add 350 µL Buffer S3, immediately invert the tube 4–6 times until a flocculent precipitate appears. Centrifuge at 12,000 × g for 10 min.

DNA Binding and Washing 5. Transfer the clear supernatant to a Spin Column (placed in a Collection Tube). Centrifuge at 12,000 × g for 60 s, discard the flow-through. 6. Add 500 µL Buffer W1, centrifuge at 12,000 × g for 60 s, discard the flow-through. 7. Add 750 µL Buffer W2, centrifuge at 12,000 × g for 60 s, discard the flow-through. 8. Centrifuge again at 12,000 × g for 1 min to remove residual ethanol.

Elution 9. Place the Spin Column into a new 1.5 mL centrifuge tube. Add 60–80 µL Elution Buffer (or ddH₂O) to the membrane. Incubate at room temperature for 1 min. 10. Centrifuge at 12,000 × g for 1 min to elute the DNA.

Ligation and Transformation Protocol (One Step Cloning)

Introduction

One Step Cloning (OSC) is a molecular cloning technique that allows seamless assembly of DNA fragments and vectors in a single reaction. Unlike traditional restriction enzyme and ligation methods, OSC uses homologous recombination or recombinase-based reactions to join DNA fragments with overlapping sequences. This approach eliminates the need for multiple digestion and ligation steps, making the cloning process faster, more efficient, and less error-prone.

Procedures

A. Ligation Reaction

1. Mix insert DNA fragments with linearized vector according to the experimental design. The total DNA amount per reaction should be 200–400 ng.

2. Prepare the following reaction mixture on ice (total volume = 10 µL):

   • 2× ClonExpress Mix (Vazyme CE): 5 µL
   • Insert DNA: X µL
   • Linearized vector: Y µL
   • ddH₂O: to 10 µL

3. Mix gently and incubate at 50 °C for 5 min.

4. The ligation product can be used immediately for transformation or stored at −20 °C for short term.

B. Transformation of Ligation Products (OSC)

1. Thaw the required competent cells on ice.
2. Add 5–10 µL of ligation product into 100 µL competent cells, mix gently, and incubate on ice for 30 min.
3. Heat shock at 42 °C for 90 s, then immediately cool on ice for 2–3 min.
4. Add 900 µL SOC medium (or LB medium without antibiotics) and incubate at 37 °C, 200 rpm for 1 h.
5. Centrifuge at 5000 rpm for 5 min, discard 900 µL of the supernatant, leaving ~100 µL. Resuspend gently.
6. Spread the suspension on LB agar plates containing the appropriate antibiotic.
7. Incubate plates at 37 °C overnight (12–16 h).

Genomic DNA Extraction Protocol (for Bacteria)

Introduction

Genomic DNA extraction is a fundamental procedure in microbiology and molecular biology, enabling the isolation of high-quality DNA suitable for downstream applications such as PCR, sequencing, cloning, and genomic analysis. In bacterial systems, the extraction process typically involves three main steps: cell lysis (to disrupt bacterial cell walls and membranes), removal of proteins and other cellular impurities, and purification of DNA using alcohol precipitation or spin column–based methods.

Procedures

A. Lysis of Gram-negative Bacteria

1. Centrifuge 1.0–5.0 × 10⁹ cells at 12,000 rpm, 2 min, discard supernatant.
2. Add 180 µL Buffer LS-2, resuspend thoroughly. Add 20 µL Proteinase K (20 mg/mL) and 10 µL RNase A (10 mg/mL), mix well. Incubate at 56 °C water bath for 10 min until the solution becomes clear.
3. Add 200 µL Buffer BS-2 and 200 µL 100% ethanol, mix well by pipetting.

B. Lysis of Gram-positive Bacteria

1. Centrifuge 0.5–2.0 × 10⁹ cells at 12,000 rpm, 2 min, discard supernatant.
2. Add 500 µL Buffer BP, resuspend pellet. Add 50 µL Lysozyme (20 mg/mL), mix thoroughly. Incubate at 37 °C water bath for 60 min (invert every 10 min).
3. Centrifuge at 12,000 rpm, 5 min, discard supernatant.
4. Add 180 µL Buffer LS-2, then add 20 µL Proteinase K (20 mg/mL) and 10 µL RNase A (10 mg/mL), mix well. Incubate at 56 °C for 10 min until transparent. If not clear, continue lysis for another 30 min, mixing every 5 min.
5. Add 200 µL Buffer BS-2 and 200 µL 100% ethanol, mix well.

C. DNA Purification (Spin Column)

1. Transfer lysate to a Bacterial DNA Mini Column. Let stand 1 min at room temperature, centrifuge 12,000 rpm, 1 min. Discard flow-through.
2. Add 500 µL Buffer WA, centrifuge 12,000 rpm, 1 min, discard flow-through.
3. Add 750 µL Buffer WB, centrifuge 12,000 rpm, 1 min, discard flow-through.
4. Repeat step 3 once.
5. Place Mini Column into a new 2 mL Collection Tube, centrifuge 12,000 rpm, 2 min to remove residual ethanol.
6. Transfer Mini Column to a clean 1.5 mL tube. Add 50 µL Elution Buffer or sterile water to the membrane center. Let stand 1 min. (Optional: heat Elution Buffer to 50–65 °C for higher yield.)
7. Centrifuge 12,000 rpm, 2 min to elute DNA. (Optional: repeat with fresh Elution Buffer for higher recovery.)

λRED Recombination (for DH5α)

Introduction

λRED Recombination is widely used in genome homologous recombination. With the help of plasmid pKD46, it can greatly reduce the length that is required for the design of left and right homologous arms, making it possible to use even 20 bp. Moreover, the replaced gene does not have to be constructed into a certain plasmid — only linear fragments with left and right homologous arms and target genes are required — which makes the entire experiment easier and cheaper.

Procedures

1. Seed the recovered bacteria DH5α [pKD46] into 5 mL of LB medium with 100 ng/mL ampicillin and shake overnight at 37 °C.
2. The next day, dilute the bacteria at a ratio of 1:25 into another 5 mL of LB medium with 100 ng/mL ampicillin. Shake at 37 °C for 1.5 h and add 10 mM L-arabinose (0.3%) to induce the λRed system.
3. Shake for another 1 h until the OD₆₀₀ is around 0.6, then cool the bacteria on ice for 10 min. Meanwhile, pre-cool ddH₂O, 10% glycerol, and centrifuge at 4 °C for 10 min.
4. Collect bacteria and wash twice with ice-cold ddH₂O in a 2 mL microcentrifuge tube (5000 rpm, 5 min).
5. Wash once with pre-cooled 10% glycerol (5000 rpm, 5 min).
6. Finally, resuspend the competent cells in 50 μL of 10% glycerol.
7. Conduct electroporation (1.8 kV/mm) and shake at 37 °C in 1 mL LB without ampicillin for 1 h.
8. Collect bacteria (8000 rpm, 3 min) and plate on LB plates with 100 ng/mL ampicillin and 100 ng/mL kanamycin, followed by incubation at 37 °C.

Preparation of Culture Media

• LB Nutrient Agar
  Qingdao Hope Bio-Technology Co., Ltd. Weigh 40.0 g of the medium, dissolve in 1000 mL distilled water, autoclave, and store for use.

• LB Broth
  Beijing Aoboxing Bio-Tech Co., Ltd. Weigh 20.0 g of the medium, dissolve in 1000 mL distilled water, autoclave, and store for use.

• TSA Agar
  Qingdao Hope Bio-Technology Co., Ltd. Weigh 40.0 g of the medium, dissolve in 1000 mL distilled water, autoclave, and store for use.

• TSB
  Qingdao Hope Bio-Technology Co., Ltd. Weigh 30.0 g of the medium, dissolve in 1000 mL distilled water, autoclave, and store for use.

• BHI
  NutriSelect® Plus Weigh 37.0 g of the medium, dissolve in 1000 mL distilled water, autoclave, and store for use.

Cell-based Experimental Protocols

Co-culture of Cancer Cells and Bacteria under Hypoxic Conditions (5% O₂, 6-well plate)

Investigation of the natural bacterial invasion efficiency in cancer cells.

Materials

Biological Samples:

• Mammalian cells (pre-adhered)
• Bacteria (cultured for 24 h)

Reagents:

• Gentamicin (15 mg/ml)
• Complete culture medium
• 1% PFA (paraformaldehyde)
• Flow buffer (PBS containing 2%FBS)
• Zombie™ APC-Cy7 Fixable Viability Dye
• PBS
• CellTrace™ Far Red Cell Proliferation Kit

Experimental Procedure

1. Preconditioning the incubator:

   Turn on the tri-gas incubator (5% O₂) at least 6 hours in advance, and place the complete culture medium inside to equilibrate.

2. Cell and bacterial preparation:

   1. Culture bacteria for 24 hours to reach the stationary phase of growth.
   2. Culture the cells until 80% confluency is achieved.

3. Invasion setup:

   1. Multiplicity of infection (MOI): Perform serial dilutions of bacterial suspensions to ensure accurate relative ratios.
   2. Co-culture duration: After 4 hours of co-incubation, bacterial internalization can be observed.

4. Bacterial CellTrace staining:

   Centrifuge the bacterial suspension and resuspend the pellet in PBS for CellTrace labeling.

5. Microscopic observation:

   Approximately 30 minutes after bacterial addition, observe and take microscopic images to document early interactions.

6. 6 - well plate setup:

   Perform all conditions in triplicate (technical replicates) and include multiple control groups.

   Ensure randomization of well placement to minimize positional bias.

   Wells containing only cells were used as negative controls to assess background signal and ensure the absence of invasion. Wells containing only culture medium were included to monitor bacterial contamination and to evaluate the efficacy of gentamicin washing.

7. Handling of HBL-100 Cells

   Special care was required when handling HBL-100 cells, as this cell line exhibits weak adherence and can be easily disrupted by PBS washing.

   During batch processing, one side of the plate was slightly elevated to create a gentle slope, and reagents were added slowly using a 1 mL pipette to avoid detachment.

   After all treatments were completed, the plates were carefully returned to a level position.

8. Gentamicin Protection Assay

   Prepare:

   • 6 mL PBS, pre-warmed in a water bath

   • 24 mL complete medium, pre-equilibrated under hypoxic conditions (5% O₂)

     (Each well of a 6-well plate contained 4 mL of medium to minimize evaporation.)

   • Gentamicin (15 mg/ml)

   The total operation time, excluding incubation, was approximately 25 minutes.

   1. Discard the culture medium (for cell-free wells, retain medium without washing—gentamicin will be added later).
   2. Wash once with 1 mL PBS to remove extracellular bacteria while maintaining cell attachment.
   3. Add 3 mL hypoxia-pre-equilibrated complete medium per well.
   4. Add gentamicin to a final concentration of 1% (30 µL per 3 mL).
   5. Incubate the plates for 30 minutes in the hypoxic incubator.

9. Cell Harvesting and Dissociation

   Prepare:

   • 2.4 mL pre-warmed Trypsin-EDTA
   • 6 mL PBS
   • 6 mL hypoxia-pre-equilibrated complete medium

   Procedure:

   1. Remove the plate from the incubator and discard the culture medium

      (for cell-free wells, directly centrifuge and discard the supernatant for viability assessment).

   2. Wash once with 1 mL PBS, then aspirate PBS completely.

   3. Add 0.4 mL Trypsin-EDTA per well and incubate until cells detach.

   4. Immediately neutralize trypsinization by adding 1 mL complete medium.

   5. Collect cell suspensions into 1.5 or 2 mL tubes, centrifuge, and remove the supernatant.

10. Optional: Live/Dead Staining

    Prepare:

    • 1.2 mL PBS (protein-free) containing Zombie APC-Cy7 diluted 1:2000 (0.6 µL stock)
    • 6 mL pre-warmed PBS

    Timing: From the end of centrifugation to completion of resuspension — approximately 11 minutes (e.g., 9:21–9:33).

    Add 0.2 mL of diluted Zombie APC-Cy7 to each 1.5 mL tube, protect from light, and incubate for 30 minutes.

    After staining, add 1 mL PBS, mix gently, centrifuge, and discard the supernatant.

11. Fixation of All Samples

    Prepare:

    • 1.2 mL pre-warmed 2% PFA
    • 0.6 mL Flow Buffer

    Resuspend each sample in 0.2 mL 2% PFA and fix at room temperature for 10–15 minutes, gently inverting the tubes during fixation.

    After fixation, centrifuge and discard the supernatant.

    Finally, resuspend in 0.1–0.2 mL Flow Buffer.

12. Use 1–2 µL of each sample for slide preparation; the remaining volume was used for flow cytometric analysis.

Protocol for Culturing Tumor Spheroids on Cell Spheroid Honeycomb Culture Chip

Materials List

Equipment

1. Flat-bottom 24-well plate
2. Cell Spheroid Honeycomb Culture Chip (VIVOID: AUT042801)

Reagents

1. Complete culture medium (DMEM + 10% FBS)
2. PBS (phosphate-buffered saline)
3. TrypLE

Seeding Procedure

1. Placement of scaffolds:

   In a biosafety cabinet, use tweezers to transfer the honeycomb scaffold into each well of the 24-well plate, ensuring the front side is facing up.

2. Pre-wetting:

   Add 1 mL culture medium onto each scaffold (avoid direct contact with the well surface), and incubate in the cell culture incubator for 15 minutes.

3. Bubble removal:

   Press the scaffold gently to the bottom of the well using a pipette tip to remove trapped air bubbles.

   Gently pipette 2–3 times over the scaffold surface to dislodge remaining bubbles.

   A dark card under the plate can help visualize trapped bubbles.

4. Cell seeding:

   Aspirate 0.5 mL medium from each well and add 0.5 mL cell suspension, gently pipetting to distribute cells evenly over the scaffold.

5. Incubation:

   Cover the plate, mark wells clearly, and place in the incubator.

   Tumor cells typically form spheroids within 1–3 days.

Medium Change

• Replace half of the medium every three days.

Spheroid Collection

1. Direct collection:

   Gently pipette the culture medium over the spheroids to resuspend them and collect the spheroid suspension.

2. Scaffold-assisted collection:

   Lift the honeycomb scaffold by its handle, invert it into a dish containing PBS, and gently tap the scaffold 2–3 times to release the spheroids into PBS. Remove the scaffold after spheroids are detached.

Matrigel-Based Tumor Spheroid Culture Protocol

I. Coating 96-Well Plates with Agarose

1. Preparation of culture medium:

   Accurately pipette 6 mL of RPMI 1640 (or DMEM) medium into two 10 mL glass serum bottles.

2. Addition of agarose:

   Add 90 mg of agarose into each bottle, seal, and place the bottles in an 80°C water bath for 30 minutes until completely dissolved.

3. Sterilization:

   After heating, sterilize the bottles at 115°C for 30 minutes in an autoclave.

4. Cooling and solidification:

   Immediately transfer the bottles to a biosafety cabinet, pour the agarose solution into a sterile reagent trough, and dispense 60 μL per well into a 96-well plate using a multichannel pipette.

   Keep the plate level for about 30 minutes to allow the agarose to solidify.

II. Preparation of Matrigel-Containing Cell Suspension

1. Cell preparation:

   Harvest MDA-MB-231 or MCF-7 cells in the logarithmic growth phase.

2. Cell counting:

   After trypsinization, count the cells and adjust the concentration to 2.0 × 10⁵ cells/mL using complete RPMI 1640 (or DMEM) medium.

3. Pre-cooling materials:

   Place an ice-filled beaker (sprayed with 70% ethanol) into the biosafety cabinet. Keep Matrigel and the culture medium on ice throughout the process.

   Note: Matrigel solidifies rapidly at room temperature—maintain all materials and pipette tips at low temperature.

4. Preparation of Matrigel mixture:

   Add 300 μL of Matrigel to 12 mL of complete medium (final concentration: 2.5% v/v) and mix quickly but gently.

   Ensure pipette tips are pre-cooled before use.

5. Preparation of final cell suspension:

   Add ~600 μL of the prepared cell suspension to the Matrigel mixture to obtain a final cell density of 1 × 10⁴ cells/mL. Mix gently and keep on ice until use.

III. Seeding Cell Suspension into Agarose-Coated 96-Well Plates

1. Seeding:

   Using a multichannel pipette, dispense 200 μL of the Matrigel–cell suspension into each agarose-coated well.

2. Centrifugation:

   Centrifuge the plate at 4°C, 1000 × g for 10 minutes using a microplate centrifuge.

   To maintain sterility, seal the plate with parafilm before centrifugation.

3. Incubation:

   After centrifugation, remove the parafilm, spray the plate with 70% ethanol, and transfer it into the cell incubator (typically 37°C, 5% CO₂).

4. Observation and medium change:

   Replace 100 μL of medium in each well on days 3, 5, and 7, and observe spheroid formation using an inverted microscope.

IV. Characterization of 3D Multicellular Tumor Spheroids

1. Observation under an inverted microscope:

   Directly place the 96-well plate under the microscope for morphological evaluation.

2. Observation under a confocal laser scanning microscope:

   Carefully collect spheroids using a pipette and wash them three times with PBS.

   Fix with 4% paraformaldehyde, stain nuclei with Hoechst 33258, wash again three times with PBS, and observe under a confocal microscope.

3. Observation under an environmental scanning electron microscope (ESEM):

   Collect spheroids as above, wash three times with PBS, fix, dehydrate, and observe under an ESEM.

Growth Factor-Induced Tumor Spheroid Formation Protocol

Serum-free medium composition:

DMEM/F12 medium supplemented with bFGF (20 ng/mL), EGF (20 ng/mL), and B27 supplement (1:50 dilution).

1. Cell Preparation

   Harvest and count the desired cell line in the logarithmic growth phase. Resuspend the cells in serum-free medium to the appropriate density.

2. Cell Seeding

   Seed 2,500 cells per well into a ULA (ultra-low attachment) flat-bottom 96-well plate, adding 250 μL of serum-free medium per well.

Ensure even cell distribution by gently pipetting the suspension up and down.

1. Culture Conditions

   Incubate the plate under hypoxic conditions (e.g., 5% O₂).

   Perform a half medium change every 3–3.5 days by carefully removing and replenishing half of the volume with fresh serum-free medium.

Primers

Notebook

Reference