Experiments

Introduction

The experimental work in our project was designed to move step by step from DNA to functional protein complexes, allowing us to test our engineered MHC Class I molecules. Each experiment builds on the previous one, forming a logical workflow that starts with molecular cloning and ends with assays measuring binding efficiency.

We began by introducing our recombinant plasmid into E. coli to produce sufficient DNA for downstream experiments. From there, we purified plasmids, expressed proteins, and validated their presence using techniques such as SDS-PAGE and BCA protein assays. To reconstitute functional MHC Class I molecules, we refolded heavy chains, β2-microglobulin, and peptides into stable complexes. Finally, we performed binding assays to compare the efficiency of wild-type and engineered variants.

Together, these experiments form the backbone of our project. They not only demonstrate our ability to take an idea from computational design to laboratory testing, but also show how synthetic biology methods can be applied systematically to address complex biological challenges. Each step highlights both technical skills and scientific reasoning, which are essential for validating our approach to improving antigen presentation in cancer immunotherapy.

Experiment 1: Transformation of Recombinant Plasmid DNA

Purpose

In experiments that use microorganisms as hosts, DNA is the most important material. To carry out molecular biology research, we need a sufficient amount of DNA. Transformation is a method that introduces foreign DNA into bacterial cells. By applying physical or chemical treatments, we make the bacterial membrane permeable so that plasmid DNA can enter the cell. Transformed bacteria can then be cultured, allowing large-scale DNA preparation.

Beyond simply obtaining DNA, transformation is widely used in the laboratory to introduce recombinant plasmids that carry new genes for research purposes.

Principle

Transformation introduces a special DNA molecule called a plasmid into host cells.
Plasmids are small, circular DNA molecules naturally found in many bacteria. They replicate independently of chromosomal DNA and often carry useful genes, such as antibiotic resistance. Researchers can insert new genes into plasmids, creating recombinant plasmids for protein expression or genetic studies.

What Cloning Means

Molecular biology cloning doesn't mean creating a complete animal. Instead, it's about making many copies of a specific piece of DNA, such as a gene. Researchers may want to study a gene or use it to make a protein. To do this, the gene is copied and placed inside a plasmid, which is a tiny, ring-shaped piece of DNA in bacteria. Plasmids are especially useful because they can carry foreign DNA and produce many copies inside bacterial cells.

To insert a gene into a plasmid, scientists use enzymes. Restriction enzymes work like molecular scissors, cutting DNA at specific sequences, while DNA ligase acts like glue, sealing the gene into the plasmid. Once this recombinant plasmid is made, it is called a "vector" because it carries the gene into bacteria. When bacteria take up the plasmid, they multiply and produce millions of gene copies.

This process is important because it allows bacteria to produce proteins for us. For example, inserting the MHC protein gene into bacteria means the bacteria can generate the protein in large amounts. This same strategy is also used to make insulin and other medicines. DNA cloning in this way gives scientists the ability to study genes, make proteins, and even design new drugs.

Competent Cells and Transformation

To make bacteria receptive to DNA, we prepare competent cells. This is usually done with E. coli grown to log phase, the stage where the bacteria are actively dividing. At this point, their surface receptors make them more likely to take up DNA.

One commonly used strain is E. coli DH5-α. After growing to log phase, cells are washed and suspended in calcium chloride. Calcium ions neutralize the negatively charged cell membrane, helping the negatively charged DNA bind to the cell surface.

Next, we use a heat shock step: rapidly heating the cell--DNA mixture, then cooling it on ice. This creates a temporary pore in the bacterial membrane, allowing plasmid DNA to enter. After recovery in nutrient-rich medium, the transformed cells express the plasmid's genes.

By plating the cells on agar containing antibiotics (such as kanamycin), only bacteria that successfully took up the plasmid (and thus the resistance gene) will grow into colonies. Each colony originates from a single transformed cell, making it a genetic clone.

Materials

• 37 °C shaking incubator
• 42 °C water bath
• Ice and ice bucket
• 1.5 mL microtubes
• Sterile spreader
• LB medium and LB agar plates (with kanamycin)
• Competent E. coli DH5-α
• Recombinant plasmid DNA (NUSA_pET28a)

Procedure

1. Keep competent cells on ice.
2. Prepare LB agar plates with kanamycin.
3. Mix 5 μL plasmid DNA with competent cells by tapping the tube (do not pipette).
4. Incubate on ice for 20 min.
5. Heat shock: 42 °C for 90 sec, then immediately place on ice for 2--3 min.
6. Add 1 mL LB medium, incubate 1 h at 37 °C shaking incubator.
7. Centrifuge briefly, remove most supernatant, leave pellet with small medium.
8. Spread onto kanamycin agar plate.
9. Incubate overnight, then observe colonies next day.

Experiment 2: Extraction and Purification of Plasmid DNA

Purpose

To extract and purify plasmid DNA from transformed E. coli. Purified plasmid DNA is essential for downstream applications such as sequencing, cloning, and protein expression.

Principle

• Why purify plasmid DNA? It's widely used for recombinant DNA, gene cloning, and protein expression.
• Amplification in E. coli: Plasmids replicate in bacteria, producing large DNA yields.
• Alkaline lysis method:
  • Lyse bacterial cells with alkaline detergent.
  • Both chromosomal and plasmid DNA denature.
  • Neutralization step → plasmids reanneal (small, circular DNA), chromosomal DNA cannot.
  • Centrifugation removes chromosomal DNA and debris.
  • Plasmids bind to filter membrane → washed → eluted in buffer.

Materials

• Plasmid purification kit (Enzynomics)
• Microcentrifuge

Procedure

1. Collect bacterial culture.
2. Lyse cells with alkaline buffer.
3. Neutralize to allow plasmid reannealing.
4. Centrifuge to pellet debris and chromosomal DNA.
5. Transfer supernatant to spin column.
6. Wash with washing buffer.
7. Elute purified plasmid DNA into new tube.
8. Store at --20 °C.

Experiment 3: Protein Expression

Purpose

To express recombinant proteins in E. coli using IPTG induction.

Principle

• IPTG Induction:
  • IPTG mimics lactose and activates the lac operon.
  • This leads to T7 RNA polymerase production, which drives transcription from the T7 promoter on the plasmid.
  • Stable induction = overexpression of target protein.
• Growth Monitoring:
  • Optical density (OD600) indicates bacterial growth phase.
  • Induction is optimal at mid-log phase (OD ~0.5--0.7).

Materials

• 37 °C shaking incubator
• LB medium (with kanamycin)
• 1 M IPTG
• Spectrophotometer

Procedure

1. Inoculate a colony into LB broth + kanamycin; incubate overnight.
2. Transfer 1--2% of pre-culture into fresh 50 mL LB medium.
3. Measure OD600 hourly.
4. At OD 0.5--0.7, add IPTG to 0.5 mM.
5. Incubate 4 h further.
6. Harvest by centrifugation.
7. Discard supernatant, freeze pellet at --20 °C.

Experiment 4: SDS-PAGE Gel Electrophoresis

Purpose

To separate proteins based on molecular weight using SDS-PAGE. By observing protein bands on the gel, we can check the expression level and approximate size of the target protein.

Principle

What is SDS-PAGE?

Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), in simple terms, is a laboratory technique for analyzing proteins based on their size. The process begins with SDS, which is a detergent that denatures proteins and gives proteins negative charges. This treatment makes proteins lose their original shapes and structures, and makes them linear. Also, SDS coats proteins with a negative charge that is proportional to individual protein's length. As a result, the separation that occurs in the gel depends mostly on protein size rather than shape or charge.

The proteins are then loaded into a polyacrylamide gel and subjected to an electric current. Since the proteins are all negatively charged, they move toward the positive electrode. The gel itself acts as a molecular filter: smaller proteins move faster and farther, while larger proteins move slower and remain closer to the top. After electrophoresis, the gel is treated with stain, which binds to proteins and makes the separated bands visible. Each band represents proteins of a specific size, allowing researchers to check protein expression and compare purification results. Overall, SDS-PAGE enables scientists to visualize and analyze proteins in a clear and convenient way.

• SDS-PAGE = Sodium Dodecyl Sulfate -- Polyacrylamide Gel Electrophoresis.
• Acrylamide gels are denser than agarose gels, making them ideal for separating proteins.
• SDS coats proteins with negative charges and denatures them into linear chains.
• When voltage is applied, proteins migrate toward the positive electrode, separated by size:
  • Small proteins → move faster (downward).
  • Large proteins → move slower (remain higher).

SDS (Detergent)

• SDS disrupts secondary, tertiary, and quaternary protein structures, unfolding them into linear chains.
• It also gives proteins a uniform negative charge so that separation depends only on size, not charge or shape.

Polyacrylamide Gel Structure

• Stacking gel (upper layer): compresses all proteins into a sharp starting band.
• Separating gel (lower layer): separates proteins by molecular weight.
• Glycine, Cl⁻ ions, and proteins migrate differently, creating conditions that align proteins in the stacking gel before separation.

Materials

• Protein sample (10 μL)
• 2× Laemmli sample buffer (Tris-HCl, SDS, glycerol, bromophenol blue, β-mercaptoethanol/DTT)
• SDS-PAGE gel kit
• SDS running buffer (Tris-Cl, glycine, SDS)
• Staining and destaining solutions

Procedure

1. Prepare SDS-PAGE gel using the kit.
2. Mix protein sample with 2× Laemmli buffer.
3. Heat mixture at 95°C for 5 minutes to denature proteins.
4. Load samples into wells and run at 180 V for ~1 hour.
5. Remove gel and stain with staining solution for 15 minutes.
6. Destain for ~30 minutes until background is clear.
7. Observe protein bands and take a photo.

Experiment 5: BCA Protein Assay

Purpose

To measure total protein concentration of samples.

Principle

• Proteins reduce Cu²⁺ → Cu⁺ in alkaline solution.
• Cu⁺ reacts with BCA to form purple complex (λ = 562 nm).
• Absorbance intensity ∝ protein concentration.
• Requires standard curve with BSA.

• Note: BCA measures total protein, not whether the proteins are correctly folded or functional.

Materials

• BSA standard
• BCA assay kit
• 96-well plate and plate reader (562 nm)

Procedure

1. Prepare BSA standards across concentration range.
2. Mix Reagent A and B = Working Reagent (50:1).
3. Add 25 μL sample + 200 μL WR per well.
4. Incubate 30 min at 37 °C.
5. Read OD at 562 nm.
6. Plot standard curve, calculate unknown concentrations.

Experiment 6: Refolding of MHC/Peptide Complexes

Purpose

To refold recombinant MHC class I heavy chain (HC) and β2m with tumor-specific peptide into functional complexes.

Principle

• HC and β2m expressed in E. coli are denatured in urea.
• Refolding buffer with L-arginine and redox system prevents aggregation.
• Order of addition: Peptide + β2m first, then HC (added gradually).
• Low temperature and slow stirring enable correct folding.

Materials

• Refolding buffer (Tris, L-arginine, glutathione redox system)
• Guanidine solution
• Protease inhibitors (PMSF, pepstatin, leupeptin)
• Recombinant HC, β2m, peptide

Procedure

1. Prepare HC (18.6 mg, ~3 μM) and β2m (13.2 mg, ~6 μM).
2. Cool 200 mL refolding buffer to 10 °C. Add inhibitors.
3. Dissolve peptide (12 mg) in buffer.
4. Dilute proteins with guanidine solution.
5. Inject β2m, then HC slowly near stir bar.
6. Incubate 8 h at 10 °C with stirring.
7. Add second aliquots → incubate 6--12 h.
8. Add final aliquots → incubate 24 h.
9. Store at 4 °C or proceed to next step.

Experiment 7: Antigen Binding Affinity Assay

Purpose

To compare binding efficiency of wild-type vs. engineered MHC--peptide complexes using a fluorescence-based assay.

Principle

• Fluorescence: molecules absorb excitation light, emit longer-wavelength light (Stokes shift).
• Intensity ∝ number of bound complexes.
• Enables comparison between different complexes.

Materials

• Nickel-coated 96-well plate
• MHC--peptide complex solutions (WT, mutant variants)
• Refolding buffer
• Microplate reader (fluorescence mode)

Procedure

1. Equilibrate wells with buffer for 5 min.
2. Load complexes (WT, mutants) + blanks.
3. Incubate 2 h to overnight at RT.
4. Wash wells 5× with buffer.
5. Add 100 μL buffer; read fluorescence.
6. Compare fluorescence intensities.