Experiments

Overview

This protocol describes the transformation of plasmids into E. coli DH5α electrocompetent cells using electroporation. The transformed cells are then recovered and plated on selective media for subsequent cryopreservation and downstream applications.

Materials

• Electrocompetent cells: E. coli DH5α
• Plasmid DNA: iGEM parts (or other plasmids of interest)
• Electroporator
• Electroporation cuvettes
• SOC medium (pre-warmed to room temperature)
• Nutrient agar plates with appropriate selection antibiotic (Kanamycin for this experiment)
• Pipettes and sterile tips: P1000, P200, P20, and P10
• Sterile 1.5 mL microcentrifuge tubes
• Sterile PCR tubes
• Sterile L-shaped spreaders (hockey sticks)
• Ice bucket with ice
• Incubator shaker (37°C)
• 95% ethanol
• Sterile distilled water

Protocol

Overview

Restriction digestion is used to cut plasmid DNA at specific recognition sites using restriction enzymes, preparing DNA for cloning, ligation, or analytical applications.

Materials

• Plasmid DNA
• PCR tubes (0.2 mL or 0.5 mL)
• Micropipettes and sterile tips
• Ice bucket

Reagents & Solutions

• 10X NEBuffer (rCutSmart® Buffer, NEB)
• Restriction enzymes (EcoRI, XbaI, NotI, SpeI, and PstI are some of the enzymes we use in the lab)
• Nuclease-free water

Protocol

Thaw reagents

1. Thaw buffer and water at room temperature. Keep restriction enzymes on ice at all times.

Reaction setup

Typical reaction volume: 10 µL (can be scaled).

Component	Volume (example 10 µL reaction)
Plasmid DNA	100–500 ng (≈1–7 µL, depending on concentration)
10X rCutSmart Buffer	1 µL
Restriction enzyme(s)	0.5–1 µL each (typically 1–10 units per reaction)
Nuclease-free water	To final volume (adjust so total = 10 µL)

Add components in the following order (to reduce enzyme inactivation):

2. Add nuclease-free water first.
3. Add 10X rCutSmart Buffer.
4. Add plasmid DNA.
5. Add restriction enzyme(s) last.

Mixing

6. Gently flick the tube or pipette up and down. Do not vortex.

Incubation

7. Incubate at 37°C for 1–2 hours.
8. For high-fidelity (HF) enzymes, 30–60 min is usually sufficient.

Optional

9. Heat-inactivate enzymes if recommended (65°C for 20 min, depending on enzyme).
10. Proceed to downstream applications (ligation, gel electrophoresis, etc.).

Overview

This protocol describes the isolation of plasmid DNA from E. coli overnight cultures using the alkaline lysis method, followed by precipitation and purification. The resulting DNA can be used for cloning, digestion, or storage.

Materials

• Micropipettes (P1000, P200, P10) with sterile tips
• Sterile 1.5 mL microcentrifuge tubes
• Bacterial culture (overnight, grown with appropriate antibiotic)
• Centrifuge capable of 13,000 rpm

Reagents & Solutions

• Solution I (buffer):
  - 50 mM glucose
  - 25 mM Tris-Cl, pH 8.0
  - 10 mM EDTA
  Add 4 mg/mL lysozyme before use.
• Solution II (Lysis):
  - 0.2 N NaOH
  - 1% (w/v) SDS
  Prepare this solution immediately before use.
• Solution III (DNA precipitation):
  - 29.5 mL glacial acetic acid
  - KOH pellets added until pH 4.8
  - Add water up to 100 mL
  - Store at room temperature, do not sterilize.
• Chloroform
• Isopropanol 100%
• Ethanol 80%
• 1X TE buffer (10 mM Tris, 1 mM EDTA, pH 8.0)

Protocol

Culture preparation

1. Inoculate the desired colony into 3–6 mL of nutrient broth with the appropriate antibiotic.
2. Incubate overnight (~16–24 h) at 37°C with shaking.

Cell collection

3. Transfer 1.5 mL of culture into a microcentrifuge tube.
4. Centrifuge at 13,000 rpm for 2 minutes to pellet cells.
5. Discard the supernatant.
6. Repeat with another 1.5 mL of culture into the same tube to increase yield.

Resuspension

7. Resuspend the pellet in 100 µL of Solution I (cold, with lysozyme).
8. Incubate at room temperature for 5 minutes to allow cell wall digestion.

Lysis

9. Add 200 µL of freshly prepared Solution II (cold).
10. Mix gently by inverting (do not vortex).
11. Incubate on ice for 5 minutes.
12. Note: Solution should become viscous/clear—indicates lysis.

Neutralization

13. Add 150 µL of Solution III.
14. Mix immediately by gentle inversion or brief vortex.
15. Incubate on ice for 5 minutes.
16. Centrifuge at 13,000 rpm for 5 minutes.
17. Carefully transfer the clear supernatant to a new tube.

Chloroform extraction

18. Add 200 µL of chloroform to the supernatant.
19. Vortex thoroughly, centrifuge 1 min at 13,000 rpm.
20. Carefully collect the aqueous (upper) phase into a fresh tube.

DNA precipitation

21. Add an equal volume of 100% isopropanol.
22. Mix by inversion and incubate at room temperature for 5 minutes.
23. Centrifuge at 13,000 rpm for 5 minutes.
24. Discard supernatant carefully.

DNA washing

25. Wash pellet with 200 µL of cold 80% ethanol.
26. Centrifuge 1 min at 13,000 rpm.
27. Discard supernatant completely.

Drying and resuspension

28. Air dry pellet no longer than 5 minutes (avoid overdrying).
29. Optional: quick spin to collect residual liquid.
30. Resuspend pellet in 50 µL of 1X TE buffer (or nuclease-free water).

Storage

31. Store plasmid DNA at –20 °C.

Overview

This protocol describes purification of DNA fragments from low-melting-point agarose gels using β-Agarase I (New England Biolabs). The method ensures recovery of intact DNA for downstream applications such as cloning, ligation, or sequencing.

Materials

• Water bath (set to 65°C)
• Heat block (set to 42°C)
• Microcentrifuge
• Sterile centrifuge tubes (1.5 mL)
• Micropipettes (P1000, P200, P10, P2) with sterile tips
• Ice bucket with ice
• Clean scalpel or razor blade

Reagents

• Low melting point agarose
• 1X β-Agarase I Reaction Buffer (NEB)
• β-Agarase I enzyme (NEB)
• NaCl (0.5 M) or alternative salts
• 100% isopropanol
• 70% ethanol (cold)
• TE buffer or nuclease-free water

Protocol

Excise the DNA band

1. Use a clean scalpel or razor blade to cut the DNA-containing band from the gel.
2. Minimize excess agarose around the DNA band.

Equilibration (standard method)

3. Wash the solid gel slice twice with 2 volumes of 1X β-Agarase I buffer on ice, 30 minutes each wash.
4. Discard the buffer after each wash.

Melting

5. Incubate the gel slice at 65°C in water bath for 10 minutes until fully molten.

Enzyme digestion

6. Cool the molten agarose to 42°C in heat block.
7. Add 1 unit of β-Agarase I per 200 µL of 1% agarose.
8. Incubate at 42°C for 1 hour.

Salt adjustment

9. Add salt to achieve one of the following concentrations for precipitation: 0.5 M NaCl, 0.3 M NaOAc, 2.5 M NH₄OAc, or 0.8 M LiCl.

Pre-clear sample

10. Chill on ice for 15 minutes.
11. Centrifuge at 14,000 RPM for 17 minutes to pellet undigested carbohydrates.
12. Carefully transfer the DNA-containing supernatant to a new tube.

DNA precipitation

13. Add 0.7 volumes of 100% isopropanol.
14. Mix thoroughly and chill on ice.
15. Centrifuge at 14,000 RPM for 17 minutes.

Washing

16. Remove supernatant carefully.
17. Wash pellet with cold 70% ethanol.
18. Centrifuge 1–2 minutes and discard supernatant.

Drying

19. Air dry the pellet at room temperature (avoid overdrying).

Resuspension

20. Resuspend DNA pellet in TE buffer or nuclease-free water.
21. Store at –20°C.

Overview

Agarose gel electrophoresis is used to verify DNA size, analyze genetic material, and check the quality/purity of DNA samples. DNA fragments migrate through agarose in an electric field, and are visualized with a nucleic acid stain. Gel electrophoresis is also an important step for DNA insert separation.

Materials

• DNA sample
• NEB 1kb ladder
• Electrophoresis chamber
• NEB 6x Loading dye
• Ethidium bromide (EtBr)
• TE buffer
• 1X TAE running buffer and for gel preparation
• Agarose
• Microwave
• Power supply
• UV transilluminator
• PCR tubes or parafilm

Protocol

Prepare agarose gel

1. Weigh 0.3 g agarose and dissolve in 30 mL of 1X TAE buffer.
2. Heat in a microwave until fully dissolved (avoid boiling over).
3. Cool to 60°C, then pour gel into casting tray with comb in place. Allow to solidify (~20–30 min).

Set up electrophoresis chamber

4. Place gel into chamber and cover completely with 1X TAE running buffer.
5. Carefully remove comb.

Prepare samples

6. Mix DNA with loading dye (this can be done in PCR tubes or on top of a clean piece of parafilm).
7. Load wells with samples: NEB 1kb ladder in one lane and DNA samples in the rest (volume depends on concentration, usually 5–10 µL).

Run gel

8. Attach electrodes (DNA runs toward the red/positive electrode).
9. Run at 90 V for 65 minutes.

Visualization

10. Soak gel in EtBr solution (0.5 µg/mL) for 15 min, then rinse in water for 10–15 min.
11. Visualize under UV or blue-light transilluminator.

Safety note: Ethidium bromide is mutagenic — handle with PPE and dispose properly or use a safer stain alternative.

Overview

DNA inserts with compatible sticky ends are ligated into the plasmid vector using T4 DNA Ligase, following the Ligation Protocol with T4 DNA Ligase by New England Biolabs.

Materials

• T4 DNA Ligase
• T4 DNA Ligase Buffer (10X)
• Vector DNA
• Insert DNA
• Nuclease-free water
• Microcentrifuge tubes
• Ice bucket

Protocol

Reaction setup

1. Prepare the ligation mixture on ice as follows (example: 20 µL total reaction, using a 1:3 vector-to-insert molar ratio):
• X µL Vector DNA
• X µL Insert DNA
• 2 µL 10X T4 DNA Ligase Buffer (thaw and resuspend at room temperature before use)
• 1 µL T4 DNA Ligase (added last)
• Add nuclease-free water to final volume

Note: NEB’s ligation calculator or standard molar ratio guidelines can be used to adjust volumes based on DNA sizes and concentrations.

Mix

2. Gently pipette up and down to mix.
3. Briefly centrifuge to collect contents at the bottom of the tube.

Incubation

4. Incubate at 16 °C overnight for maximum efficiency.

Heat inactivation

5. Inactivate the enzyme at 65 °C for 10 minutes.

Transformation

6. Chill the reaction on ice.
7. Transform 1–5 µL of the ligation mixture into 20 µL of competent cells.

Overview

This protocol describes the purification of plasmid DNA from E. coli using the ZymoPURE™ Plasmid Miniprep Kit with the centrifugation method. The kit allows rapid recovery of up to 100 µg of high-quality plasmid DNA suitable for downstream applications including cloning, sequencing, and transfection.

Materials

• Bacterial culture: 0.5–5 mL overnight E. coli culture (LB + antibiotic)
• ZymoPURE™ Plasmid Miniprep Kit reagents:
• P1 (Red)
• P2 (Blue)
• P3 (Yellow)
• Binding Buffer
• Wash 1
• Wash 2 (with ethanol added, if required)
• Elution Buffer
• Zymo-Spin™ II-PX Columns with collection tubes
• 1.5 mL sterile microcentrifuge tubes
• Microcentrifuge (≥ 10,000 RPM)
• P1000, P200, and P10 micropipettes and sterile tips

Protocol

Preparation

1. Add the required ethanol to ZymoPURE™ Wash 2 concentrate before use.
2. Equilibrate all reagents to room temperature.
3. Ensure P2 and Binding Buffer are fully dissolved (warm to 30–37°C if precipitated).

Cell Lysis & Neutralization

4. Pellet 0.5–5 mL of overnight bacterial culture at full speed (15–20 sec). Discard supernatant.
5. Resuspend pellet in 250 µL ZymoPURE™ P1 (Red) by pipetting or vortexing.
6. Add 250 µL ZymoPURE™ P2 (Blue). Gently invert 8–10 times (do not vortex). Incubate 3 min.
7. Solution should appear clear, purple, and viscous.
8. Add 250 µL ZymoPURE™ P3 (Yellow). Invert until solution turns yellow and precipitate forms.
9. Centrifuge 6.5 min at 14,000 RPM.

DNA Binding

10. Transfer 600 µL supernatant to a new 1.5 mL tube.
11. Add 260 µL Binding Buffer, vortex 15 sec.

Column Purification (Centrifugation)

12. Transfer entire lysate–binding buffer mix to a Zymo-Spin™ II-PX Column in a collection tube.
13. Incubate 1 min, then centrifuge ≥ 10,000 RPM for 1 min. Discard flow-through.
14. Add 800 µL Wash 1. Centrifuge ≥ 10,000 RPM for 1 min. Discard flow-through.
15. Add 800 µL Wash 2. Centrifuge ≥ 10,000 RPM for 1 min. Discard flow-through.
16. Add 200 µL Wash 2. Centrifuge ≥ 10,000 RPM for 1 min. Discard flow-through.
17. Centrifuge column again ≥ 10,000 RPM for 1 min to remove residual wash buffer.

Elution

18. Place column into a clean 1.5 mL tube.
19. Add 25 µL Elution Buffer directly to the column matrix. Incubate 2 min.
20. Centrifuge ≥ 10,000 × g for 1 min to elute plasmid DNA.
21. Store DNA at ≤ –20°C.

Overview

This protocol describes the preparation of a sterile, accurately quantified 5.00 mM stock solution of RDX (MW = 222.12 g/mol) in acetone at a final volume of 25.00 mL, suitable for experimental use, following institutional safety procedures.

Calculations

Desired concentration × volume = moles required

\[ 5.00 \, \text{mM} = 0.00500 \, \text{mol/L} \]

\[ \text{Volume} = 25.00 \, \text{mL} = 0.02500 \, \text{L} \]

Moles = concentration × volume

\[ \text{moles} = 0.00500 \, \text{mol/L} \times 0.02500 \, \text{L} \]

\[ \text{moles} = 0.000125000 \, \text{mol} \]

Convert moles to mass using MW of RDX (222.12 g/mol)

\[ \text{mass (g)} = \text{moles} \times \text{MW} \]

\[ \text{mass} = 0.000125000 \, \text{mol} \times 222.12 \, \text{g/mol} \]

\[ 222.12 \, \text{g/mol} \times 0.000125000 \, \text{mol} = 0.0277650 \, \text{g} \]

Conversion

\[ 0.0277650 \, \text{g} \times 1000 \, \text{mg/g} = 27.7650 \, \text{mg} \]

Materials & Equipment

• Analytical balance (readability ≤0.1 mg)
• Amber glass vial or amber glass bottle with tight cap (volume ≥ 50 mL) — acetone compatible
• Volumetric flask and pipettes to make up to 25.00 mL
• Acetone (reagent grade)
• Weighing paper or small weighing boat
• Forceps/tweezers (optional)
• Labels and marker (chemical name, concentration, solvent, date, preparer, hazard info)
• Fume hood

Protocol

Pre-label container

1. Label volumetric flask: “RDX — 5.00 mM in acetone; 25.00 mL; prepared [date]; prepared by [name]; hazard: [type of hazard]”

Weigh compound

2. Tare weighing boat on the analytical balance. Add RDX until the balance reads 27.77 mg. Use tweezers/clean spatula; avoid touching with bare hands. Immediately transfer the weighed solid into the pre-labeled volumetric flask or vial in the fume hood.

Add solvent to dissolve

3. In the hood, add approximately 15–20 mL of acetone to the flask/vial to dissolve the solid. (If using a 25.00 mL volumetric flask: add solvent to dissolve then bring to the mark — see next step.) Swirl gently or invert to mix; do not vortex if using open glassware. Do not heat the acetone (it is flammable). If dissolution is slow, gentle swirling is sufficient.

Bring to final volume

4. After the compound is completely dissolved, add acetone to bring the solution to the 25.00 mL mark (final volume). If using a graduated cylinder, transfer quantitatively into a 25.00 mL volumetric flask and then adjust to the mark with acetone. Ensure meniscus at calibration line at eye level.

Mix & transfer

5. Cap the flask/vial and invert several times to ensure homogeneity. If using a volumetric flask, invert ~10–15 times.

Storage

6. Store at 4°C at the recommended temperature. Include hazard label on storage container.

Waste & decontamination

7. Dispose of all acetone rinses, contaminated wipes, and weighing boats in the designated organic solvent waste container, following institutional hazardous waste procedures. Do not pour acetone or compound into sink. Decontaminate balance area and hood per lab SOP.

Overview

Assess whether E. coli detects RDX when grown in defined medium with antibiotic selection by comparing biological responses across multiple exposure levels and appropriate controls.

Materials

• Sterile culture tubes (suitable volume for 6 mL working volume) and caps
• Sterile pipette tips (sized for planned liquid handling)
• Sterile serological/transfer pipettes
• Sterile inoculating tools
• Chemical fume hood (for handling RDX)
• Incubator and shaker
• Micropipettes (P10, P20, P200, P1000)
• Sterile test tubes

Reagents

• M9 defined medium (prepared, sterile)
• Kanamycin stock (1000× concentrated) — label with concentration, lot, date, preparer
• Authorized E. coli strain culture (per your lab’s IBC approval)
• RDX stock (amber vial, 5.00 mM in acetone, 25 mL) — properly labeled and with SDS on hand
• Acetone (reagent grade) — if needed for vehicle controls or rinses

Protocol

Preparation of test tubes

1. Pour 6 mL of previously prepared and sterilized M9 defined medium into desired amount of test tubes.
2. Make sure appropriate aseptic techniques are being employed (work close to Bunsen burner).
3. Add 6 µL of 1000× concentrated kanamycin antibiotic.

Inoculation

4. With a sterile inoculation loop, transfer pre-incubated bacteria into test tubes to ensure the same transfer amount.

Adding RDX

5. *Note: This step was carried out in the chemistry department under supervision.
6. Transfer calculated volumes of RDX 5 mM stock into test tubes.

Target [RDX] (µM)	For 6 mL LB Tubes: Volume from 5 mM Stock (µL)	For 6 mL LB Tubes: Volume from 10 mM Stock (µL)
0.00	0	0
0.10	0.1	24
0.30	0.4	72
0.44	0.50	105.2
0.50	0.60	120
1.00	1.20	240
2.00	2.40	480
3.80	4.60	912
4.40	5.30	1056
5.00	6.00	1200
6.20	7.40	1488

Incubation

6. Incubate at 37°C at 160 RPM for 48 hours. Make sure to take note of any observations seen at 24 and 48 hours.

Disposal

7. Dispose of medium with RDX in the waste container and follow the laboratories disposal protocols.