1 Preparation of Experimental Reagents and Media

Each group will prepare the corresponding volume/quantity of reagents/media.

1.1 Experimental Objective

To prepare reagents needed for subsequent experiments.

Genetic Engineering:

Protein Engineering:

1.2 Experimental Procedure: Preparation of Reagents for Genetic Engineering

1.2.1 Preparation and Sterilization of Solutions

LB Medium

A total of 300 ml is needed, but 400 ml will be prepared to ensure sufficient volume during aliquoting.

Preparation Method: [Tryptone: 4g, Yeast Extract: 2g, NaCl: 4g] + [400 ml pure water], stir well in a beaker, aliquot 20 ml into 10 conical flasks, and the remainder into a blue-cap bottle, and autoclave (loosen the cap for the blue-cap bottle).

LB Solid Medium (1.5%)

[Tryptone: 2g, Yeast Extract: 1g, NaCl: 4g] + [200 ml pure water] + [Agar powder: 3g], prepare directly in a 500 ml conical flask, do not stir, cover, and autoclave (loosen the cap for the blue-cap bottle).

Calcium Chloride (for competent cell preparation)

[0.55g CaCl2] + [50 ml pure water], prepare directly in a 100 ml blue-cap bottle, and autoclave.

Plastic Consumables

1.5 ml centrifuge tubes, 20 pcs, packaged in a plastic bag, and autoclaved.

1.2.2 Pouring Plates

Add Ampicillin (1 ml + 1 μl, final concentration 50 μg/ml) to the sterilized and cooled LB solid medium below 60°C, mix well, and pour plates into 90 mm plastic petri dishes, approximately 10-15 ml per plate, at least 10 plates needed.

1.3 Experimental Procedure: Preparation of Reagents for Protein Engineering

Prepare solutions according to the formulas for Solution A and Solution B, adjust pH, filter through a membrane, and degas by ultrasound. Each group prepares 1L of Solution A and 500 ml of Solution B.

Filter and degas 1L of deionized water by ultrasound, prepare 1L per group.

Prepare 20% ethanol solution, filter through a membrane, and degas by ultrasound.

2 Inoculation (Preparation for Plasmid Extraction)

Add 15 ml of autoclaved LB medium from 1.2.1 to a sterile 50 ml centrifuge tube, add 15 μl of Ampicillin stock solution (1 ml + 1 μl, final concentration 50 μg/ml), mix well. Add 150 μl of DH5α bacterial culture.

Cap and remove from the biosafety cabinet, incubate in a shaker at 220 rpm, 37°C, overnight.

3 Fundamental Experiments in Genetic Engineering/Molecular Biology

3.1 Objective

The molecular biology/genetic engineering experiments on the 11th are conducted individually. Each student must perform the basic molecular biology/genetic engineering experiments. The experiments on the 12th will be conducted in groups.

3.2 Experimental Procedures

3.2.1 Plasmid Miniprep and Concentration Measurement

3.2.2 PCR

3.2.2.1 Fragment Preparation

PCR Conditions for Fragment

Estimated Time: 1 hour (Morning Session)


PCR Conditions for pet28a Fragment

Estimated Time: 1 hour (Noon Break)

3.2.2.2 Purification of PCR Products

Due to the complexity of the PCR system, which contains templates, primers, enzymes, and other impurities that may hinder seamless cloning, purification is necessary to obtain the target gene fragment.

After purification, measure DNA concentration again and calculate the yield of the purification process.

3.2.3 Agarose Gel Electrophoresis

Prepare the gel during the PCR amplification of the fragment. In the afternoon, add the loading buffer and then run the gel electrophoresis.

(1) Prepare the gel (do this during the morning PCR session; due to the limited number of gel trays, prepare two small gels per group).

Agarose Gel Electrophoresis

Our target gene and vector backbone are approximately 700 bp and 6000 bp, respectively, so prepare a 1% agarose gel.

Dissolve 1 g of agarose in 100 ml TAE buffer (not pure water!) in a 250 ml Erlenmeyer flask, cover with foil, and heat in a microwave until dissolved. Cool to approximately 60°C, add nucleic acid dye, mix, and pour into the gel tray. Cover with foil and allow to solidify in the dark.

（2）Preparation of Electrophoresis Samples

Mix a small amount of the purified PCR product with loading buffer (6X), in a 1:5 ratio.

Suggested: Take 10 µl each of the fragments in a 0.2 ml EP tube, add 2-3 µl loading buffer, and mix well.

（3）Loading Samples

（4）Electrophoresis and Analysis

Run at 90V. After verifying the molecular weight, store the samples at -20°C for use on the 12th.

4 Competent Cells, Seamless Cloning, and Transformation

4.1 Experimental Procedures

4.1.1 Preparation of Competent Cells

Upon arriving at the laboratory, first, pre-cool the calcium chloride solution prepared and sterilized on the 10th in an ice bath.

Transfer 1.25 ml of the bacterial culture into 1.5 ml centrifuge tubes (distributed into 8 tubes) and place them on ice for 10 minutes to cool to 0°C. Centrifuge the bacterial culture at 4°C, 4000 rpm for 10 minutes, and discard the supernatant (invert the tubes to drain the medium thoroughly).

Add 150 μL of ice-cold 0.1 M CaCl₂ to resuspend, combine two tubes into one, and place on ice for 30 minutes (resulting in 4 tubes total). Centrifuge again at 4°C, 4000 rpm for 10 minutes, discard the supernatant, and add 100 μL of ice-cold 0.1 M CaCl₂ to resuspend to obtain BL21 competent cells for immediate transformation. If not used immediately, add an equal volume of 50% glycerol, rapidly freeze in liquid nitrogen, and store at -80°C.

4.1.2 Seamless Cloning

Design the seamless cloning system according to the manual (classroom exercise; concentrations vary per individual, so each person's table will differ).

08/Experimental Procedure

08-1/Recombinant Reaction

1. Concentration Determination of Linearized Vectors and Insert Fragments

If the linearized vector and insert fragments have been purified through a gel recovery kit, and the absorbance reading shows no apparent bands or smears remaining, you can use Nanodrop or similar spectrophotometers to measure concentration. However, the A260/A280 ratio should only be considered reliable between 1.8 and 2.0. Instruments like Nanodrop, Qubit, and PicoGreen are recommended for concentration measurement. When sample concentrations are below 10 ng/μl, different instrument models may yield significantly varying readings based on A260.

2. Calculation of Vector and Insert Fragment Usage:

For single-fragment source recombination, the minimum recommended vector usage is 0.03 pmol, and the minimum recommended insert fragment usage is 0.06 pmol (with a vector-to-insert molar ratio of 1:2).

For multi-fragment source recombination, the minimum DNA usage for each fragment is 0.03 pmol (with a vector-to-insert molar ratio of 1:1).

The amount of DNA used in these calculations can be approximated using the following formulas:

3. Prepare the Following Reaction Systems on Ice:

  1. X/Y is calculated based on the calculated vector and insert fragment amounts. To ensure amplification accuracy, vectors and insert fragments should be concentrated before reaction setup and used in no less than 1 μl volumes.
  2. Positive Controls-1 and -2 employ plasmids as templates, with amplification lengths <3 kb. Primer Tm values should be >60°C.
  3. Positive Control-3 uses genomic DNA as a template. Use high-fidelity polymerases for amplification.
  4. Fragment length, GC content, and sequence complexity affect reaction time. High GC content in insert fragments can hinder the reaction; consider adding DMSO or betaine to enhance amplification.

4. Gently mix using a pipette (avoid vortex mixing) and briefly centrifuge the reaction mixture to the bottom of the collection tube.

5. Recombination Reaction Conditions:

Single-Fragment Recombination: Incubate at 50°C for 5 minutes; cool to 4°C or place on ice immediately.

2-3 Fragment Recombination: Incubate at 50°C for 15 minutes; cool to 4°C or place on ice immediately.

4-5 Fragment Recombination: Incubate at 50°C for 30 minutes; cool to 4°C or place on ice immediately.

4.1.3 Transformation

Recombinant Product Transformation

  1. Thaw chemically competent cells on ice (e.g., Fast-T1 Competent Cell, Vazyme #C505).
  2. Add 5-10 μl of the recombinant product to 100 μl of competent cells. Gently flick the tube to mix (avoid vortexing) and incubate on ice for 30 minutes.
    • The volume of recombinant product should not exceed 1/10 of the volume of competent cells used.
  3. Heat shock the cells at 42°C for 30 seconds, then immediately place them back on ice for 2-3 minutes.
  4. Add 900 μl of SOC or LB medium (without antibiotics) and incubate at 37°C with shaking for 1 hour (rotation speed 200-250 rpm).
  5. Pre-warm the required LB agar plates containing the appropriate antibiotic at 37°C in an incubator.
  6. Centrifuge at 5,000 rpm (2,500 x g) for 5 minutes and discard 900 μl of the supernatant. Resuspend the bacterial pellet in the remaining medium and spread it gently on the pre-warmed agar plate with the antibiotic.
  7. Invert the agar plates and incubate at 37°C for 12-16 hours.

Each experimental group will have four sets of samples (AC, AD, BC, BD), with each set plated on 1-2 LB plates and incubated overnight at 37°C.

5 Biotechnology Visit and Fermentation Inoculation

5.1 Fermentation (Inoculation)

Upon arrival in the morning, distribute the fermentation seed culture prepared on the 13th into the remaining three 250 ml sterilized culture bottles from step 2.2, adding at least 25 ml of seed culture to each (using sterile Pasteur pipettes). Incubate at 37°C, 220 rpm for 5-10 hours.

Afternoon: Visit to the biotechnology facility.

5.2 Fermentation (Induction)

After the visit in the afternoon, return to the laboratory and add 125 μl of IPTG stock solution to each bottle (the stock is 1 M, with a final concentration of 1 mM). Incubate overnight at 37°C, 220 rpm for induction.

6 Bacterial Collection, Ultrasonic Disruption, Separation of E. coli Lysate, Protein Electrophoresis

6.1 Collection and Resuspension

Fill the fermentation broth into 50 ml centrifuge tubes, not exceeding 40 ml per tube. Balance the tubes, and centrifuge multiple times, ensuring each group collects bacterial cells from at least 200 ml of fermentation broth. Centrifuge at 4000 x g for 10 minutes. Resuspend the collected bacterial pellet in solution A, at a ratio of 1 g of cells to 10 ml of solution A. Transfer 1 ml of resuspended cell culture to a new 1.5 ml EP tube for protein electrophoresis.

6.2 Ultrasonic Disruption

Disrupt the resuspended bacterial cells using ultrasonic waves. Ensure that the centrifuge tubes containing the cells are placed in an ice bath within a beaker. Use ultrasonic settings of 5 seconds on, 5 seconds off, for a total of 20 minutes.

6.3 Centrifugal Separation of E. coli Lysate

6.3.1 Experiment Objective

Our target protein is located in the E. coli lysate supernatant. However, cell disruption produces many insoluble impurities (e.g., cell wall debris), so the first purification step is centrifugation to separate the insoluble precipitate from the soluble protein solution.

6.3.2 Experimental Procedures

Centrifuge the E. coli lysate obtained above using 1.5 ml EP tubes, with 1 ml per tube, at 4°C, 17,000 x g for 40 minutes. After centrifugation, transfer the supernatant to a new centrifuge tube, and resuspend the precipitate with 1 ml of solution.

6.4 Protein Electrophoresis

6.4.1 Experiment Objective

To further confirm the expression status of our protein (the proportion of target protein in the lysate supernatant, solubility of the target protein), protein electrophoresis is required.

6.4.2 Experimental Procedures

Sample Preparation

For three 0.2 ml EP tubes, take 100 μl each of resuspended cell culture, lysate supernatant, and lysate precipitate. Add at least 20 μl of 6× protein loading buffer to each tube, and incubate in a PCR thermocycler for 20 minutes.

Loading

Use pre-cast gels. Each person should practice loading at least two wells. Each group should load samples that include the three prepared in 4.2.1 and a protein marker.

Electrophoresis

Run at 160V for 40 minutes.

Staining

Incubate the gel in staining solution on a shaker for 20 minutes to visualize bands.

7 Cell Culture and Passaging Protocol

Materials Needed

Steps

  1. Preparation:

    • Ensure that all materials and reagents are sterile.

    • Warm the cell culture media, PBS, and trypsin-EDTA to 37°C in the incubator.

  2. Observing Cells:

    • Check the cells under an inverted microscope to assess their confluency. Ideally, cells should be 70-80% confluent for passaging.

  3. Removing Old Media:

    • Gently aspirate the old culture media from the flask or plate without disturbing the cell layer.

  4. Washing Cells:

    • Add 5-10 mL of PBS to wash the cells. Swirl gently and then aspirate the PBS. This step removes residual media and dead cells.

  5. Adding Trypsin-EDTA:

    • Add enough trypsin-EDTA solution to cover the cell layer (typically 1-2 mL for a T25 flask).

    • Incubate at 37°C for 2-5 minutes. Monitor the cells under the microscope. Cells should start to round up and detach.

  6. Stopping Trypsin Activity:

    • Once most cells are detached, add an equal volume of fresh complete cell culture media (containing serum) to neutralize the trypsin.

    • Gently pipette up and down to dislodge any remaining attached cells.

  7. Collecting Cells:

    • Transfer the cell suspension to a sterile centrifuge tube.

    • If needed, count the cells using a hemocytometer or automated cell counter.

  8. Dilution and Re-seeding:

    • Dilute the cell suspension to the desired cell concentration with fresh media.

    • Transfer the diluted cell suspension to new tissue culture flasks or plates (e.g., 1:5 or 1:10 dilution for passaging).

  9. Incubation:

    • Place the newly seeded flasks or plates in the incubator set at 37°C with 5% CO₂.

    • Allow cells to adhere and grow for 24-48 hours before the next observation.

  10. Documentation:

    • Record the date, cell passage number, and any observations regarding cell morphology and health.

Notes

  1. Always use aseptic techniques to prevent contamination.

  2. Adjust the passaging frequency based on the specific growth characteristics of your cell line (e.g., every 2-4 days).

  3. Dispose of waste properly in biohazard containers.

  4. This protocol should guide you through the cell culture and passaging process efficiently. Let me know if you need any additional information!

8 Tube Formation Assay Protocol

Materials Needed

Steps

  1. Preparation:

    • Thaw Matrigel overnight at 4°C. Keep it on ice until use.

    • Warm complete endothelial cell media to 37°C.

  2. Cell Culture:

    • Culture HUVECs in complete media until they reach 70-80% confluency.

    • Wash cells with PBS and detach using trypsin-EDTA. Neutralize trypsin with complete media.

  3. Cell Counting:

    • Count the cells using a hemocytometer or automated cell counter.

    • Adjust the cell density to approximately 1 × 10⁶ cells/mL in complete media.

  4. Matrigel Coating:

    • Add 50-100 µL of Matrigel per well in a 96-well plate. Swirl gently to ensure even distribution.

    • Allow the Matrigel to polymerize by incubating at 37°C for 30-60 minutes.

  5. Seeding Cells:

    • Add 100-200 µL of the HUVEC suspension (1 × 10⁶ cells/mL) onto the polymerized Matrigel.

    • Gently swirl the plate to distribute the cells evenly.

  6. Incubation:

    • Place the plate in the incubator (37°C, 5% CO₂) and allow the cells to adhere and form tubes for 4-6 hours or overnight, depending on the experimental design.

  7. Treatment (Optional):

    • If testing the effects of treatments (e.g., CBD MMPs-VEGF), add the treatments to the cells during this incubation period.

  8. Observation:

    • After incubation, observe the plate under an inverted microscope.

    • Capture images of the tube-like structures formed by the endothelial cells.

  9. Analysis:

    • Analyze the tube formation using quantitative parameters such as total tube length, number of junctions, and branching points. Use image analysis software if available.

  10. Documentation:

    • Record your observations, including any differences between control and treatment groups, and save images for further analysis.

Notes

  1. Ensure all materials and tools are sterile to prevent contamination.

  2. Repeat the experiment in triplicates or more for statistical validity.

  3. Adjust cell density or incubation time based on specific experimental requirements.

  4. This protocol will guide you through conducting a Tube Formation Assay effectively. Let me know if you need any further details!

9 Alkaline Phosphatase (ALP) Staining Protocol

Materials Needed

Steps

  1. Cell Culture and Differentiation:

    • Culture MC3T3-E1 cells (or your cell line) in complete growth media.

    • Induce osteogenic differentiation by switching to osteogenic media, if applicable, and continue culturing for 7-14 days (early differentiation stages).

  2. Fixing Cells:

    • After the desired differentiation period, aspirate the media from the cells.

    • Wash the cells twice with PBS to remove any remaining media.

    • Fix the cells by adding 4% paraformaldehyde (PFA) and incubating for 10-15 minutes at room temperature.

    • Wash cells again with PBS to remove excess PFA.

  3. Staining Procedure:

    • Prepare the ALP staining solution according to the manufacturer’s instructions (e.g., BCIP/NBT substrate solution).

    • Add the ALP staining solution to the fixed cells, covering the entire cell layer.

    • Incubate at 37°C for 30-60 minutes or until the desired level of blue/purple color development is achieved.

  4. Washing and Observation:

    • Once staining is complete, aspirate the staining solution and wash the cells gently with PBS.

    • Observe the stained cells under a microscope. ALP-positive cells will show blue/purple staining, indicating ALP activity.

  5. Documentation:

    • Take images of the stained cells for further analysis and quantification.

  6. Optional:

    • If desired, measure the staining intensity by solubilizing the dye and quantifying the absorbance using a spectrophotometer.