Procedure

1. Pellet 1–5 mL overnight culture by centrifugation >8000 rpm (≈6800 × g) for 3 min at room temperature (15–25 °C).
2. Aspirate supernatant and resuspend pellet in 250 µL Buffer P1; transfer to a microcentrifuge tube.
3. Add 250 µL Buffer P2; invert 4–6× until clear (≤5 min).
4. Add 350 µL Buffer N3; invert 4–6×.
5. Centrifuge 10 min at 13,000 rpm (≈17,900 × g).
6. Load 800 µL supernatant onto a QIAprep column; spin 30–60 s; discard flow-through.
7. Wash with 500 µL Buffer PB; spin 30–60 s; discard.
8. Wash with 750 µL Buffer PE; spin 30–60 s; discard. Spin again 1 min to dry.
9. Place column in a clean 1.5 mL tube; add 25 µL nuclease-free H₂O to membrane center; wait 1 min; spin 1 min to elute.

Procedure

1. Set thermocycler to 98 °C.
2. Find T_m for each template.
3. Prepare PCR mix in a microcentrifuge tube.

4. Aliquot 25 µL PCR MM into tubes.
5. Prepare a negative control (replace template with water).
6. Store PCR samples at 4 °C as needed.

Thermocycler Settings (25 µL Reaction)

Step	Temperature (°C)	Time
Initial Denaturation	98	30 s
35 Cycles	98	10 s
	60	30 s
	72	90 s
Final Extension	72	5 min
Hold	4	—

* Extension times ~30 s/500 bp; optimize per insert.

Post-PCR Checks

1. Run 0.8–1% agarose gel; visualize bands.
2. If single correct band: PCR cleanup → quantify.
3. If multiple bands: gel extract the correct band → quantify.
4. If no/faint band: try annealing gradient; repeat at 50 µL.

Procedure

1. Add 5× Buffer PB to 1× PCR reaction; invert 4–6×.
   • If color turns orange/violet, add 10 µL 3 M sodium acetate (pH 5.0) — should return to yellow.
2. Load to column; spin 1 min; discard flow-through.
3. Wash with 750 µL Buffer PE; spin 1 min; discard.
4. Spin again 1 min to dry.
5. Elute with 20–30 µL EB/water (wait 1 min before spin).

Procedure

1. Master mix for 30 µL (per reaction):
   • 25 µL 2× polymerase master mix
   • 2.5 µL 10 µM forward primer
   • 2.5 µL 10 µM reverse primer
2. In tube: 30 µL nuclease-free water + 30 µL 2× mix (final 60 µL).
3. Touch a colony with sterile toothpick; swirl into tube.

Program

Step	Temperature (°C)	Time
Initial Denaturation	98	30 s
35 Cycles	98	10 s
	60	30 s
	72	90 s
Final Extension	72	5 min
Hold	4	∞

Proceed to gel electrophoresis as needed.

Procedure

1. Weigh agarose per expected fragment size.

2. Microwave until dissolved; cool to 45–50 °C.
3. Add Sybr Safe (e.g., 5 µL/50 mL); swirl.
4. Pour gel; let solidify; remove comb and place gel in chamber.
5. Cover with 1× TAE. Load 1 kB ladder (2 µL).
6. Mix sample + loading dye on parafilm; load. Avoid bubbles.
7. Run 75–120 V until dye front ~80% down gel.
8. Image under UV/blue light.

Procedure

1. Prepare a 20 µL digestion mix:
   • Calculate vector volume from concentration (e.g., to get 1 µg from 50 ng/µL, use 20 µL).
   • Add water to 20 µL total volume with 2 µL 10× buffer and 1 µL each enzyme.
2. Mix gently; quick spin.
3. Incubate 37 °C for 5–15 min; heat-inactivate if applicable.

Reaction Setup

Prepare a 20 µL reaction:

* ~1 µg of a 3 kb plasmid ≈ 1 pmol ends.

** If enzymes cannot be heat-inactivated, purify DNA prior to ligation.

Procedure

1. Incubate 37 °C for 10 min.
2. Heat-inactivate 80 °C, 2 min.

Procedure

1. For a 20 µL ligation (1:3 vector:insert molar ratio):
   • 2 µL 10× T4 Ligase Buffer*
   • 50 ng vector (≈0.020 pmol)
   • 37.5 ng insert (≈0.060 pmol)
   • Nuclease-free H₂O to 19 µL
   • 1 µL T4 DNA Ligase (add last)

* Thaw/resuspend buffer at room temp (do not hand-warm).

2. Mix gently; quick spin (avoid bubbles).

Thermocycler Program — Cohesive (Sticky) Ends

Step	Temperature (°C)	Time
Cycle	10 ↔ 30	30 s each · 71 cycles
Final	65	10 min
Hold	10	∞

3. Heat-inactivate at 65 °C for 10 min.
4. Chill on ice and transform 10 µL into 100 µL competent cells.

Method

The electrometric method of Wilbur and Anderson (1948) in which the time required (in seconds) for a saturated CO2 solution to lower the pH of 0.012 M Tris⋅HCl buffer from 8.3 to 6.3 at 0°C is determined. The time without enzyme is recorded at T0; with enzyme, T.

Reagents

• 20 mM Tris⋅HCl buffer, pH 8.3. Store in an ice bath at 0-4°C before and during use.
• Carbon dioxide saturated water. Bubble CO2 gas through 200 ml ice cold water for 30 minutes prior to assay using sodastream (pressing button for 5 seconds for high carbonation). During saturation process, store water at 0-4°C in an ice bath.
• Enzyme

Add 1mL of cell lysate of E. coli with recombinant carbonic anhydrase to the initial volume of Tris.

Procedure

Blank Determination: Add 3.0 ml of chilled 20 mM Tris⋅HCl buffer, pH 8.3 to a 50 ml falcon tube in ice. Maintain temperature at 0-4°C and record pH.
Add 2 ml of chilled CO2 saturated water to the Tris buffer. Immediately start a stop watch and record the time required for the pH to drop from 8.3 to 6.3. Record this time as T0.

Enzyme Determination: Add 2.0 ml of chilled 20 mM Tris⋅HCl buffer, pH 8.3 to a 50 ml falcon tube on ice. Maintain temperature at 0-4°C and record pH. Add 1 ml of enzyme sample and allow for pH to equilibrate. Quickly add 2 ml of CO2 saturated water and record the time required for the pH to drop from 8.3 to 6.3. Record this time as T.

Calculations:

https://www.worthington-biochem.com/products/carbonic-anhydrase/assay

MeCN (or DCM/THF) → Cold MeOH/EtOH

Lysate Cells and Weigh -
Lyse cells (via sonication or an alternative method) and weigh 10 grams of the resulting cell lysate–Poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHBV) granule mixture.

Dissolve (5–10% w/v) -
Add 100 mL acetonitrile to a beaker (for 10g PHBV)
Add 10 grams of lysate + plastic granule mix; stir at RT until fully dissolved (30–120 min).
Gentle warming to 30–35 °C can speed up; do not exceed solvent bp. Acetonitrile boils at 170ish, you can pump up the temp to much greater than 35 just a couple tens of degrees lower than the boiling point. keep in mind the melting point of pla as well, you don’t want to exceed that.

Prepare Nonsolvent Bath -
In a chilled beaker with a stir bar, add 500 mL of cold methanol. Place in an ice bath; aim for ~4–10 °C.

Precipitate (Slow Addition) -
With vigorous stirring, pour the PHBV solution slowly into the cold methanol (not the reverse). A white flocculent precipitate forms immediately. Rinse the solution vessel with a few mL of MeCN and add dropwise to maximize yield.

Collect & Wash - Vacuum filter through a Büchner funnel. Wash cake with 2× 50 mL cold MeOH to strip residual solvent.

Dry - Air-dry the cake in the hood 30–60 min, then vacuum-dry at 30–40 °C to constant mass. Gently crumble any soft agglomerates with a spatula to obtain free-flowing powder.

Source: team wiki notes. :contentReference[oaicite:0]{index=0}

1) Make 5% βME Laemmli buffer (per sample, 20 µL)
10 µL 2× Laemmli (no βME)
9 µL H₂O
1 µL β-mercaptoethanol (βME) → 5% v/v
For n samples, make (n+1)× master mix (20 µL per sample).

2) Denature samples (final 1× Laemmli)
Thaw samples briefly at 23 °C, then place on ice.
Mix 20 µL 5% βME Laemmli + 20 µL protein sample (lysate supernatant).
Cap tightly; quick vortex + spin.
Heat 95 °C, 10 min.
Spin 1000 g, 5 min. Keep on ice.

3) Set up gel/tank (4–20% SDS–PAGE)
Remove bottom tape; place gel in holder.
If single gel, insert back plate.
Fill inner chamber with 1× TGS; fill outer tank to ~¼ height.
Remove comb straight up.
Rinse wells with ~15 µL 1× TGS each.

4) Load & run
Load 5 µL prestained ladder.
Load 10 µL denatured supernatant per sample.
Run 100 V, 10 min (stacking).
Then 130 V until dye front ~⅔ down gel (~50–60 min).
Do not run off the gel.

5) Retrieve gel
Open cassette; transfer gel to a staining container.

6) Fairbanks staining (A → B → C → D; 100–150 mL each)
For each solution (A, then B, then C, then D):
Submerge gel in 100–150 mL solution.
Microwave 80–90 s until just boiling.
Immediately move to fume hood; rock gently 1 min to cool.
Drain and rinse gel with ddH₂O.

7) Image
Place gel on white screen/plate in the imager.
Select Coomassie Blue program; adjust sliders for clarity.
Save images (e.g., PNG/JPG) and archive a .tif copy.

Fairbanks recipes (1 L each)
A: 0.05% Coomassie (50 mL of 1%), 25% isopropanol (250 mL), 10% acetic acid (100 mL), ddH₂O to 1 L (~600 mL).
B: 0.005% Coomassie (5 mL of 1%), 10% isopropanol (100 mL), 10% acetic acid (100 mL), ddH₂O to 1 L (~795 mL).
C: 0.002% Coomassie (2 mL of 1%), 10% acetic acid (100 mL), ddH₂O to 1 L (~898 mL).
D: 10% acetic acid (100 mL), ddH₂O to 1 L (~900 mL).

Source: team wiki notes. :contentReference[oaicite:1]{index=1}

A) Prep: Reagent & Standards

Equilibrate & mix reagent
Gently invert the Coomassie Plus (Bradford) Assay Reagent a few times (do not shake). Bring to room temp before use.

Prepare protein standards (BSA recommended)
Use BSA 2.0 mg/mL stock to make a standard curve spanning your working range.

Standard working ranges:
Plate / test-tube (standard): 100–1500 μg/mL (linear with BSA ~125–1000 μg/mL).
Low-range (micro format): 1–25 μg/mL.

Example dilutions are provided below (use the same diluent as your samples).

Example BSA standards (standard range, for plate or test tube)
2000, 1500, 1000, 750, 500, 250, 125, 25, 0 μg/mL

Example BSA standards (low-range, for micro format)
25, 20, 15, 10, 5, 2.5, 0 μg/mL

Add 50 μL standard or sample to labeled tubes.
Add 1.5 mL Bradford reagent, mix well.
Zero spectrophotometer with water; read A595.
Blank-correct and quantify from the standard curve (linear range with BSA ~125–1000 μg/mL).

ThermoFisher user guide (for reference): MAN0011344