Escherichia coli Culture ( E. coli )

Materials

• Media: LB (routine), SOC (post-transformation recovery), TB/Terrific Broth (high density).
• Antibiotics (working): Amp 100 µg/mL; Kan 50 µg/mL; Cm 25 µg/mL; Spec 50 µg/mL.
• Plates: LB-agar + appropriate antibiotic.
• Consumables: Sterile loops/tips, 14/50 mL tubes, baffled flasks (5x culture volume).
• Equipment: 37 °C incubator, shaking incubator (200-250 rpm, 25 mm orbit), spectrophotometer (OD600).

Media (1 L)

• LB: 10 g tryptone, 5 g yeast extract, 10 g NaCl; dHO1 L; autoclave.
• LB-Agar: LB + 15 g agar; autoclave, cool to 55 °C, add antibiotic, pour plates.
• SOC (ready-made preferred) for post-transformation.
• TB: 12 g tryptone, 24 g yeast extract, 4 mL glycerol; dHO900 mL; autoclave. After cooling add 100 mL 10x TB salts (0.17 M KHPO, 0.72 M KHPO).

Protocols

Day 0: Streak for Single Colonies

1. Warm selective plate (room temp, lid down).
2. Flame loop touch glycerol stock/colony streak for isolation (quadrant streak).
3. Invert and incubate 12-16 h at 37 °C (or 24 h if slow).

Day 1: Starter Culture (Overnight)

1. Pick one well-isolated colony inoculate 3-5 mL LB + antibiotic in a 14 mL tube.
2. Incubate 37 °C, 220-250 rpm, ~12-16 h (loosely capped for aeration).

• Expression strains for toxic proteins: consider 30 °C overnight to reduce stress.

Day 2: Main Culture

1. Inoculate baffled flask with 1:100 from starter (e.g., 5 mL into 500 mL LB/TB + antibiotic).
2. Incubate 37 °C, 200-250 rpm until desired OD600:
   • Mid-log: 0.4-0.6 (ideal for induction/transformation-related assays).
   • Late-log: 0.8-1.0 (yield vs. stress trade-off).
3. Proceed to the downstream step (induction, plasmid prep, harvest, etc.).

Protein Expression (BL21(DE3))

• At OD600 0.5-0.7, add IPTG 0.1-1 mM (typical 0.5 mM).
• Express 2-4 h at 30-37 °C (fast, may form inclusion bodies) or 16-20 h at 16-20 °C (solubility).
• Harvest: 4,000-6,000xg, 10 min, 4 °C; proceed to lysis.

Source: Addgene protocol

Protocol

1. Take competent cells out of -80 °C and thaw on ice (approximately 20-30 mins).
2. Remove agar plates (containing the appropriate antibiotic) from storage at 4 °C and let them warm up to room temperature and then (optional) incubate in a 37 °C incubator.
3. Mix 1-5 µL of DNA (usually 10 pg-100 ng) into 20-50 µL of competent cells in a microcentrifuge or falcon tube. Gently mix by flicking the bottom of the tube with your finger a few times.
   Note: Transformation efficiencies will be approximately 10-fold lower for ligation of inserts to vectors than for an intact control plasmid.
4. Incubate the competent cell/DNA mixture on ice for 20-30 mins.
5. Heat shock each transformation tube by placing the bottom 1/2 to 2/3 of the tube into a 42 °C water bath for 30-60 secs (45 secs is usually ideal, but this varies depending on the competent cells you are using).
6. Put the tubes back on ice for 2 min.
7. Add 250-1,000 µL LB or SOC media (without antibiotic) to the bacteria and grow in a 37 °C shaking incubator for 45 min.
   Note: This outgrowth step allows the bacteria time to generate the antibiotic resistance proteins encoded in the plasmid backbone so that they will be able to grow once plated on the antibiotic containing agar plate. This step is not critical for Ampicillin resistance but is much more important for other antibiotic resistances.
8. Plate some or all of the transformation onto a 10 cm LB agar plate containing the appropriate antibiotic.
9. Incubate plates at 37 °C overnight.

Summary

20uL of sample was mixed with 6.6uL of 4X LDS-PAGE loading (Cat# NP0008) buffer containing
10% v/v beta-mercaptoethanol. Samples were boiled for 10min at 95C and then spun in a
centrifuge at 20,000xg for 30seconds. The entire 26.6uL volume was loaded into wells of a
self-cast 4% stacking/10% resolving gel (1.5mm thickness). The gel was run at 200V for ~40min
in a tris-glycine buffer according to Bio-Rad instructions for SDS-PAGE (Laemmli) buffer system.

Source : Biorad Mini-PROTEAN 3 Cell Protocol

Protocol

Gel Cassette Preparation

1. Ensure glass plates, casting frames, and stands are clean and dry.
2. Place Casting Frame upright with cams open.
3. Insert Spacer Plate (label side up) with a Short Plate in front.
4. Slide plates into frame with short plate facing forward; lock cams to seal.
5. Place the cassette on the casting stand, making sure the gasket seals.
6. Repeat for second gel if needed.

Gel Casting (Hand-Cast)

Discontinuous Gel (standard for SDS-PAGE)

1. Insert comb, mark gel at 1 cm below comb teeth, remove comb.
2. Prepare resolving gel mix (all but APS/TEMED); degas 15 min.
3. Add APS/TEMED, pour to mark, overlay with water or t-amyl alcohol, polymerize 45-60
   min, rinse.
4. (Optional overnight storage: overlay with 5 mL 1x resolving gel buffer.)
5. Prepare stacking gel mix, degas 15 min.
6. Dry area above resolving gel, add APS/TEMED, pour to top of short plate.
7. Insert comb carefully, polymerize 30-45 min, remove comb, rinse wells.

Continuous Gel

1. Prepare and degas monomer solution.
2. Add APS/TEMED, pour to top of short plate.
3. Insert comb, polymerize 45-60 min, remove comb, rinse wells.

Ready-Gel Option

1. Remove precast gel from pouch, take out comb, rinse wells.
2. Cut a dotted line at the bottom and peel sealing tape to expose the gel edge.

Module Assembly

1. Remove gel sandwiches from the casting stand; unlock from frames.
2. Insert into electrode assembly with short plates facing inward.
3. Close clamps to seal; lower inner chamber into tank.
4. Add ~125 mL running buffer to inner chamber and ~200 mL to outer tank.

Sample Prep & Loading

1. Mix 20 µL protein sample with 6.6 µL 4x LDS-PAGE buffer + 10% -mercaptoethanol.
2. Boil 10 min at 95 °C, spin 20,000xg for 30 s.
3. Load full volume (~26.6 µL) into wells with gel-loading tips; load slowly to avoid
   puncturing wells.

Running the Gel

1. Place lid on tank, align color-coded leads.
2. Run at 200 V constant for ~35-40 min in Tris-glycine (Laemmli) buffer.

Gel Removal & Cleanup

1. Turn off power, disconnect leads, remove lid.
2. Pour off the buffer before opening clamps.
3. Separate sandwich plates; loosen gel with wedge tool or razor and float into fixative or
   transfer buffer.
4. Rinse tank, frames, and electrode assembly with distilled water after use.

Notes:

• Ensure proper gasket seal to prevent leaks.
• Avoid leaving alcohol overlay on gel for > 1 h to prevent dehydration.
• Refer to the Bio-Rad Mini-PROTEAN 3 Cell User Guide for full details.

Summary

The gel was transferred onto a 0.2um nitrocellulose membrane according to Bio-Rad instructions using the Mini-Transblot cell. 100V was applied for 60min in tris-glycine buffer containing 20% v/v MeOH. The blot was blocked with 1% casein in 1x PBS followed by probing with mouse anti-His at 0.1 ug/mL in 1% casein (Genscript A00186) followed by HRP-conjugated goat anti-Mouse (subclass specific) (Jackson Cat 115-035-164) at 0.16 ug/mL in 1% casein. The blot was then developed using DAB (Sigma Cat D-5905) For further details, please refer to the source.

Source: Biorad Mini Trans-Blot Electrophoretic Transfer Cell Protocol

Protocol

Prep & Safety

1. Clean before use: Wash tank, cassettes, fiber pads with mild lab cleaner; rinse DI water.
2. Safety interlock: Power off before removing lid. Max ratings (instrument level): 400 VDC, 500 W, 40 °C.
3. Cooling: Store blue cooling unit at 20 °C; insert frozen for every run. Use a stir bar in the tank (mix fast) to keep temperature/ions uniform.

Buffers & Materials

1. Transfer buffer (cold, 4 °C): Common options (see Section 3.3 of manual):
   • Tris-glycine (25 mM Tris, 192 mM glycine 20% MeOH, pH 8.3) - standard SDS-PAGE transfers.
   • 48 mM Tris/39 mM glycine 20% MeOH, pH 9.2 - alternative.
   • 10 mM NaHCO/3 mM NaCO 20% MeOH, pH 9.9 - useful for basic proteins.
   • Do not adjust pH unless specified; use analytical-grade methanol; do not reuse buffer.
2. Membrane:
   • PVDF - prewet 15 s in 100% MeOH, then equilibrate in transfer buffer.
   • Nitrocellulose - equilibrate in transfer buffer (binding often improved with 20% MeOH).
3. Cut to size: Membrane, filter papers, and fiber pads trimmed to gel; pre-soak 15-20 min in transfer buffer (also equilibrate gel).

Cassette Sandwich (Cathode Anode)

1. On cassette base (gray side down): Fiber pad Filter paper Gel (no bubbles) Membrane (no bubbles) Filter paper Fiber pad.
2. Gently roll out all bubbles (glass rod/pipette).
3. Close and latch cassette without shifting layers.
4. For acidic transfers (Section 2.3): swap gel/membrane order so membrane sits on the cathode side (proteins move toward ).

Tank Setup & Run

1. Place cassettes into electrode module (color-coded orientation), add frozen blue cooling unit.
2. Insert module into tank; fill to blotting mark with cold transfer buffer (950 mL with cooling unit). Add stir bar and mix fast.
3. Attach lid (safety interlock), connect leads.
4. Typical conditions (from Table 3.1):
   • Overnight/standard: low voltage (e.g., 30 V constant).
   • High-intensity wet transfer (mini gels): ~100 V, ~1 h (watch heating).
   • Always keep the unit cold (cooling unit + stirring).
5. On completion: power off, disconnect, remove cassette, recover membrane, rinse briefly (e.g., TBST/PBS), proceed to blocking.

Notes:

• Pre-equilibrate gels (15-20 min) to remove salts and prevent shrinkage.
• Orientation check: Gel toward cathode (), membrane toward anode (+) (except acidic mode).
• High MW proteins: consider less MeOH, add 0.05-0.1% SDS, longer time, basic buffer (pH 9.5-10); PVDF preferred.
• Low MW proteins: 0.2 µm nitrocellulose or PVDF; shorten time to avoid blow-through.
• Do not adjust buffer pH unless directed; changes can spike current/heat.
• Bubbles/contact: Most common cause of swirls/missing bands-always roll out bubbles and use fresh pads/papers

Source: Invitrogen Human IL-10 ELISA Kit - Product Information Sheet

Protocol

Note: Binding capacity depends on target and conditions; as a planning estimate, ~50 µL bead slurry often yields on the order of tens of micrograms of purified His-tagged protein. Optimize for your construct.

Buffer preparation

• 2x IMAC Buffer (0.04 M sodium phosphate, 0.6 M NaCl, pH 7.4)
• 2 M Imidazole (pH 7.4)
• Prepare native binding, wash, and elution buffers from concentrates according to the resin/system used.

Bead equilibration

1. Resuspend bead slurry thoroughly.
2. Dispense 50 µL slurry to a 1.5 mL tube; add 200 µL binding buffer, mix, place on magnet, discard supernatant.

Binding

1. Add 1.0 mL clarified lysate to equilibrated beads.
2. Incubate 30 min at room temperature with mixing or ~850 rpm shaking.
3. Place on magnet; keep supernatant as flow-through if needed.

Wash

1. Add 500 µL wash buffer, resuspend; pellet on magnet; remove supernatant.
2. Repeat wash 3x total; remove residual buffer after final wash.

Elution

1. Add 100 µL elution buffer; shake ~2 min at ~850 rpm.
2. Place on magnet and collect eluate. Optional: repeat and pool.

Source: Invitrogen Human IL-10 ELISA Kit (KHC0101) - Product Information Sheet

Kit components

Required materials not supplied

• Distilled or deionized water
• Calibrated adjustable precision pipettes and tubes for dilutions
• Beakers, flasks, and cylinders for reagent preparation
• Microtiter plate reader (450 nm; optional 550 nm reference)
• Plate washer

Protocol

Buffer preparation

• Dilute 16 mL Wash Buffer Concentrate (25x) with 384 mL water to make 1x Wash Buffer.
• Store concentrate and 1x buffer at 2-8 °C; use diluted buffer within 14 days.

Dilute standards

Note: This assay has been calibrated against the International Standard preparation (92/516) for human IL-10 (NIBSC, Hertfordshire, UK, EN6 3QG).

1. Reconstitute Hu IL-10 Standard to 5,000 pg/mL with Standard Dilution Buffer. Refer to the standard vial label for instructions. Swirl or mix gently and allow the contents to sit for 10 minutes to ensure complete reconstitution. Label as 5,000 pg/mL human IL-10. Use the standard within 1 hour of reconstitution.
2. Add 50 µL Reconstituted Standard to one tube containing 450 µL Standard Diluent Buffer and mix. Label as 500 pg/mL human IL-10.
3. Add 200 µL Standard Diluent Buffer to each of 7 tubes labeled as follows: 250, 125, 62.5, 31.2 ,15.6, 7.8, and 0 pg/mL human IL-10.
4. Make serial dilutions of the standard as shown in the following dilution diagram. Mix thoroughly between steps.
5. Discard any remaining reconstituted standard. Return the Standard Diluent Buffer to the refrigerator.

Prepare 1x Streptavidin-HRP

Note: Prepare 1X Streptavidin-HRP within 15 minutes of usage. The Streptavidin-HRP (100X) is in 50% glycerol, which is viscous. To ensure accurate dilution:

1. For each 8-well strip used in the assay, pipet 10 µL Streptavidin-HRP (100X) solution, wipe the pipette tip with clean absorbent paper to remove any excess solution, and dispense the solution into a tube containing 1 mL of Streptavidin-HRP Diluent. Mix thoroughly.
2. Return the unused Streptavidin-HRP (100X) solution to the refrigerator.

Perform Assay (Total assay time: 5 hours)

Note: Allow all components to reach room temperature before use. Mix all liquid reagents prior to use. Determine the number of 8-well strips required for the assay. Insert the strips in the frames for use. Re-bag any unused strips and frames, and store at 2°C to 8°C for future use.

1) Bind Antigen

2) Add Biotin Conjugate

1. Add 100 µL Human IL-10 Biotin Conjugate to each well (except blanks). Incubate 2 h at room temperature. Wash 4x.

3) Add Streptavidin-HRP

1. Add 100 µL of 1x Streptavidin-HRP to each well (except blanks). Incubate 30 min at room temperature. Wash 4x.

4) Add TMB

1. Add 100 µL TMB to each well. Incubate 30 min at room temperature protected from light; solution turns blue.

5) Add Stop Solution

1. Add 100 µL Stop Solution to each well; tap to mix. Solution turns yellow.

Read plate and calculate

• Read absorbance at 450 nm within 2 h (optional 550 nm reference).
• Use 4-parameter logistic (4-PL) curve fit; subtract background if available.
• Multiply sample values by dilution factor.

Performance characteristics

• Range: 0-500 pg/mL
• Sensitivity: less than 1 pg/mL
• Precision: typical intra-assay CV 3-5%; inter-assay CV 3-6%

NEB enzyme digestion (Ndel & EcoRI double digest)

Source: NEB product pages and general guidelines for Restriction Endonucleases (e.g., NdeI, EcoRI/EcoRI-HF, CutSmart Buffer)

Kit components (typical NEB catalog)

• NdeI
• EcoRI or EcoRI-HF
• CutSmart Buffer (10x)
• Gel Loading Dye (optional

Compatibility note: NdeI and EcoRI/EcoRI-HF both show 100% activity in 1x CutSmart Buffer at 37 °C, enabling a one-tube double digest.

Required materials not supplied

• DNA (plasmid or PCR product)
• Nuclease-free water
• Optional: BSA (already included in CutSmart), DNA ladder, agarose gel reagents
• Optional cleanup: spin column or magnetic-bead DNA purification kit

Notes

Unit definition

For restriction enzymes, 1 Unit (U) is the amount of enzyme required to completely digest 1 µg of DNA in 1 hour at the recommended temperature in a 50 µL reaction.

General assay/handling conditions

• Recommended working temperature: 37 °C for both NdeI and EcoRI/EcoRI-HF.
• Keep total glycerol 5% v/v in the final reaction (enzyme stocks are ~50% glycerol).
• Use 1x CutSmart as the buffer (includes BSA).
• Typical enzyme usage: 5-10 U per enzyme per µg DNA (do not exceed 10% of reaction volume with enzyme).

Methylation & star activity

• Routine Dam/Dcm methylation does not block EcoRI or NdeI.
• To minimize star activity: keep the correct buffer, avoid prolonged incubations, high glycerol, or very low ionic strength; do not over-digest.

Protocol

1. Thaw & mix. Thaw 10x CutSmart Buffer and DNA on ice. Keep enzymes on a cold rack. Briefly vortex buffer and water; do not vortex enzymes (flick or pipette gently)

2. Assemble on ice in this order:

   a) Water b) 10x CutSmart c) DNA d) NdeI e) EcoRI/EcoRI-HF.
   Mix by gentle pipetting; quick-spin to collect contents.

3. Incubate. Place at 37 °C for 30-60 min. For difficult templates or >1 µg DNA, extend to 60-90 min (avoid very long incubations to minimize star activity)

4. Check completion (optional). Run 2-5 µL on an agarose gel to verify the expected digestion pattern.

5. Heat-inactivate or clean up.

   • EcoRI-HF/EcoRI and NdeI are generally heat-inactivatable at 65 °C for ~20 min; however, because double-digests combine enzymes, the most reliable approach is column or bead cleanup to remove enzymes and buffer prior to ligation or downstream steps

6. Proceed to ligation or analysis. For cloning, purify the digested insert/vector (gel extraction if you need size selection), then set up ligation.

Glycerol Stock

Source: Addgene Protocols Creating Bacterial Glycerol Stocks for Long-term Storage of Plasmids

*Follow the steps for Inoculating an Overnight Liquid Culture.

Materials

• Glycerol
• Sterile Loop
• Overnight Bacterial Culture
• Screwtop or Cyrocrovial Tube

Protocol

1. After you have bacterial growth, add 500 µL of the overnight culture to 500 µL of 50% glycerol in a 2 mL screw top tube or cryovial and gently mix.
2. Freeze the glycerol stock tube at -80°C. The stock is now stable for years, as long as it is kept at -80°C. Subsequent freeze and thaw cycles reduce shelf life.
3. To recover bacteria from your glycerol stock, open the tube and use a sterile loop, toothpick or pipette tip to scrape some of the frozen bacteria off of the top. Do not let the glycerol stock unthaw! Streak the bacteria onto an LB agar plate.
4. Grow your bacteria overnight at the appropriate temperature. Growth conditions, including copy number and growth temperature, can be found on your plasmid's information page. The next day you will be able to start an overnight culture for plasmid DNA prep the following day

https://www.addgene.org/protocols/create-glycerol-stock/
https://www.addgene.org/protocols/create-glycerol-stock/
https://www.addgene.org/protocols/inoculate-bacterial-culture/

Polymerase Chain Reaction (PCR)

Source: Q5® High-Fidelity DNA Polymerase

Kit components

Required materials not supplied

• Deoxynucleotide (dNTP)
• Nuclease-free water

Notes

Unit Definition

• One unit is defined as the amount of enzyme that will incorporate 10 nmol of dNTP into acid insoluble material in 30 minutes at 74°C.

Unit Assay Conditions

• 25 mM TAPS-HCl (pH 9.3 @ 25°C), 50 mM KCl, 2 mM MgCl₂, 1 mM β-mercaptoethanol, 200 µM dNTPs including [³H]-dTTP and 15 nM primed M13 DNA.

Protocol

Assemble all reaction components on ice. Each component should be gently mixed before adding to the reaction in a sterile thin-walled PCR tube. The Q5 High-Fidelity DNA Polymerase may be diluted in 1X Q5 Reaction Buffer just prior to use to reduce pipetting errors. The entire reaction should be mixed again to ensure a homogeneous, consistent mixture. Collect all liquid to the bottom of the tube with a quick centrifuge spin if necessary. Overlay the sample with mineral oil if using a PCR machine without a heated lid.

Quickly transfer the reactions to a thermocycler preheated to the denaturation temperature (98°C) and begin thermocycling.

https://www.neb.com/en-us/products/m0491-q5-high-fidelity-dna-polymerase

Primer Design Guidelines:

1. Template

Use of high-quality, purified DNA templates greatly enhances the success of PCR. Recommended amounts of DNA template for a 50 µL reaction are as follows:

2. Primers

Oligonucleotide primers are generally 20–40 nucleotides in length and ideally have a GC content of 40–60%. Computer programs such as Primer3 can be used to design or analyze primers. The best results are typically seen when using each primer at a final concentration of 0.5 µM in the reaction. However, amplification of certain long, complex DNA targets (≥ 5 kb) may benefit from using a lower primer concentration (~0.2 to 0.3 µM).

3. Mg²⁺ and additives

Mg²⁺ concentration of 2.0 mM is optimal for most PCR products generated with Q5 High-Fidelity DNA Polymerase. When used at a final concentration of 1X, the Q5 Reaction Buffer provides the optimal Mg²⁺ concentration.

4. Deoxynucleotides

The final concentration of dNTPs is typically 200 µM of each deoxynucleotide. Q5 High-Fidelity DNA Polymerase cannot incorporate dUTP and is not recommended for use with uracil-containing primers or templates. Should uracil-containing primers or templates be used, we recommend Q5U® Hot Start High-Fidelity DNA Polymerase.

5. Q5 High-Fidelity DNA Polymerase concentration

We generally recommend using Q5 High-Fidelity DNA Polymerase at a final concentration of 20 units/mL (1.0 unit/50 µL reaction). However, the optimal concentration of Q5 Hot Start High-Fidelity DNA Polymerase may vary from 10–60 units/mL (0.5–3 units/50 µL reaction) depending on amplicon length and difficulty.

6. Buffers

The 5X Q5 Reaction Buffer provided with the enzyme is recommended as the first-choice buffer for robust, high-fidelity amplification. For difficult amplicons, such as GC-rich templates or those with secondary structure, the addition of the Q5 High GC Enhancer can improve reaction performance. The 5X Q5 Reaction Buffer is detergent-free and contains 2.0 mM Mg²⁺ at the final (1X) concentration.

7. Denaturation

An initial denaturation of 30 seconds at 98°C is sufficient for most amplicons from pure DNA templates. Longer denaturation times can be used (up to 3 minutes) for templates that require it.

8. Annealing

Optimal annealing temperatures for Q5 High-Fidelity DNA Polymerase tend to be higher than for other PCR polymerases. The NEB Tm Calculator should be used to determine the annealing temperature when using this enzyme. Typically, use a 10–30 second annealing step at the recommended Ta. A temperature gradient can also be used to optimize the annealing temperature for each primer pair.

9. Extension

The recommended extension temperature is 72°C. Extension times are generally 20–30 seconds per kb for complex, genomic samples, but can be reduced to 10 seconds per kb for simple templates (plasmid, E. coli, etc.) or complex templates < 1 kb. Extension time can be increased to 40 seconds per kb for cDNA or long, complex templates, if necessary.

10. Cycle number

Generally, 25–35 cycles yield sufficient product. For genomic amplicons, 30–35 cycles are recommended.

11. 2-step PCR

When primers with annealing temperatures ≥ 72°C are used, a 2-step thermocycling protocol (combining annealing and extension into one step) is possible.

12. Amplification of long products

When amplifying products > 6 kb, it is often helpful to increase the extension time to 40–50 seconds/kb.

13. PCR product

The PCR products generated using Q5 High-Fidelity DNA Polymerase have blunt ends. If cloning is the next step, then blunt-end cloning is recommended. If T/A-cloning is preferred, the DNA should be purified prior to A-addition, as Q5 High-Fidelity DNA Polymerase will degrade any overhangs generated. The Monarch® Spin PCR & DNA Cleanup Kit (5 µg) (NEB #T1130) is recommended as an efficient method for purification and concentration up to 5 µg of high-quality, double-stranded and single-stranded DNA.

Addition of an untemplated -dA can be done with Taq DNA Polymerase.

Source: Addgene protocol

Protocol

Buffer Preparation

• Add supplied Monarch RNase A to Monarch Buffer B1 to 100 µg/mL; store at 4 °C.
• Add isopropanol to Buffer BZ (0.43 volumes per volume of BZ).
• Add ethanol to Buffer WZ (4 volumes per volume of WZ).

Keep buffer bottles tightly closed.

Plasmid Miniprep Protocol (Centrifugation)

1. Pellet 1-5 mL culture (not to exceed 15 OD units) for 30 s. Discard supernatant.
2. Resuspend pellet in 200 µL Buffer B1.
3. Add 200 µL Buffer B2, invert 5-6x, incubate 1 min. Do not vortex.
4. Add 400 µL Buffer B3, invert to neutralize. Incubate 2 min.
5. Centrifuge 2-5 min.
6. Transfer supernatant to Spin Column S2D; spin 1 min. Discard flow-through.
7. Add 200 µL Buffer BZ; spin 1 min.
8. Wash with 400 µL Buffer WZ; spin 1 min.
9. Transfer column to clean tube.
10. Add 30 µL Buffer EY to matrix center; wait 1 min; spin 1 min to elute DNA.

Lactobacillus Culture

Source: Difco Lactobacilli Agar

Materials

• For Agar:
  • Dehydrated MRS Agar powder: 48 g in 1 L purified water (per manual).
  • Alternative version: 70 g/L depending on product revision.
• For Broth:
  • Same MRS formula, omit agar (38 g/L per manual).
• Purified water
• Autoclave & sterilization equipment
• Sterile glassware (flasks, bottles, pipettes)
• Sterile Petri dishes (for agar)
• Inoculating tools (loops, needles)
• Incubator set to appropriate temperature (30-37 °C)
• Control (reference) culture strain for performance testing

Protocol

Agar & Broth Preparation

1. Dissolve medium.
   • Weigh the dehydrated powder (agar or broth) and suspend in 1 L purified water.
   • Heat gently with stirring, and boil 1-3 min (per version) to completely dissolve.
2. Sterilize. Autoclave at 121 °C for 15 minutes.
3. Quality check / controls. After sterilization, test using control cultures to verify expected Lactobacillus growth.

Using the Media: Inoculation & Culture

For Plates (Solid Agar)

1. Pour ~15-20 mL of molten agar (cooled to ~45-50 °C) into sterile Petri dishes.
2. Add sample (diluted bacterial suspension) by spread/dip; e.g., 1 mL per plate when using pour-plate counts.
3. Rotate plate to distribute inoculum; let solidify 5-10 min on a flat surface.
4. Incubate: typically 35 °C for ~24 h in 5% CO₂ (or 30 °C up to 72 h) depending on strain.

For Broth

1. Inoculate MRS Broth with sample.
2. Incubate 35 °C for ~3 days, or 30 °C for ~5 days, under aerobic conditions (CO₂ enrichment optional per strain).
3. Subculture to solid medium as needed for isolation and enumeration.

Gel Electrophoresis

Equipment

• Casting tray
• Well combs
• Voltage source
• Gel box
• UV light source
• Microwave

Reagents

• 10x TAE (ThermoFisher)
• Agarose
• GelGreen Nucleic Acid Gel Stain

Protocol

Preparing TAE

Dilute 10x TAE Buffer with water for 1x TAE concentration.

Pouring a Standard X% Agarose Gel

1. Measure X g of agarose.
2. Mix agarose powder with 100 mL 1x TAE in a microwavable flask.
3. Microwave for 1-3 min until the agarose is completely dissolved (but do not overboil the solution, as some of the buffer will evaporate and thus alter the final percentage of agarose in the gel. Many people prefer to microwave in pulses, swirling the flask occasionally as the solution heats up).
4. While dissolved, add X mL of GelGreen Nucleic Acid Gel Stain (add to molten agarose at 1x final concentration).
5. Pour the agarose into a gel tray with the well comb in place, ensuring no bubbles.
6. Place newly poured gel at 4 °C for 10-15 mins OR let sit at room temperature for 20-30 mins, until it has completely solidified.

Conducting Gel Electrophoresis

1. Add a loading buffer to each of your DNA samples and mix well. Use 5 µL of loading buffer per 25 µL of sample.
2. Once solidified, place the agarose gel into the gel box (electrophoresis unit).
3. Fill the gel box with 1x TAE (or TBE) until the gel is covered.
4. Carefully load your samples into the additional wells of the gel.
5. Run the gel at 80-150 V until the dye line is approximately 75-80% of the way down the gel. A typical run time is about 1-1.5 hours, depending on the gel concentration and voltage.
6. Turn OFF power, disconnect the electrodes from the power source, and then carefully remove the gel from the gel box.
7. Using any device that has UV light, visualize your DNA fragments. The fragments of DNA are usually referred to as bands due to their appearance on the gel.

Protocol

Buffer & Reagents (keep everything cold, 2–8 °C)

• Lysis buffer (typical, cytosolic proteins): 50 mM Tris-HCl pH 7.5–8.0, 150 mM NaCl, 5–10% glycerol
  Optional: 1 mM DTT (or 2 mM β-ME), 1 mM MgCl₂, 0.1–1 mM EDTA*
• Protease inhibitor: 1 tablet/10–50 mL (EDTA-free if you need metalloproteins).
• DNase I: 5–10 µg/mL (add with MgCl₂) to reduce viscosity post-lysis.
• Clarification supplies: ice bucket, 0.45 µm prefilter (optional), centrifuge bottles.

*If you will purify a His-tagged metalloprotein, avoid EDTA.

Prep & Harvest

1. Grow culture to OD₆₀₀ ~0.6–1.0 (or as required for expression). Chill culture on ice 10–15 min to slow proteolysis.
2. Pellet: 4,000–6,000 ×g, 10–15 min, 4 °C.
3. Resuspend pellet in cold lysis buffer at 3–5 mL per gram wet pellet. Add protease inhibitors. Keep on ice.

Set Up Disruptor

• Clean/flush the system (water → 20–30% ethanol → water) if needed.
• Prime with a small volume of lysis buffer to remove air.
• Cooling: Use built-in heat exchanger, inline chiller, or coil tubing in an ice bath on the outlet. Aim to keep outlet <10 °C.

Operating Parameters (typical for E. coli)

• Pressure: 10–20 kpsi (~70–140 MPa). Start at 10–12 kpsi; increase if lysis is poor.
• Passes: 1–3 passes. Most E. coli lyse >90% by 2 passes.
• Flow rate: As per instrument (e.g., 30–120 mL/min). Avoid cavitation; maintain steady back-pressure.
• Viscosity control: Add DNase I to the collection vessel at start of run (final 5–10 µg/mL + 1–5 mM MgCl₂).

Lysis Procedure

1. Prefilter the resuspension through 0.45 µm to catch clumps that can clog valves.
2. Load the cold cell slurry into the feed reservoir.
3. Run Pass 1 at 10–12 kpsi; collect into a pre-chilled bottle containing DNase I/MgCl₂. Gently swirl to mix.
4. Assess quickly: take 50–100 µL for either:
   • Phase-contrast check (ghost cells vs intact), or
   • Quick OD drop/viscosity check with pipetting feel.
5. Run Pass 2 (and Pass 3 if needed) at 12–18 kpsi. Keep outlet cold. Avoid foaming.
6. Hold on ice 10 min to let DNase act and viscosity drop.

Clarification

1. Centrifuge 15,000–20,000 ×g, 20–30 min, 4 °C to remove debris.
2. Save supernatant (crude lysate). For very fine debris, polish with 0.45 µm → 0.22 µm filtration if downstream requires.
3. Snap-sample for SDS-PAGE (pre/post lysis) to document efficiency.