How it works

Molarity is moles of solute per litre of solution. Multiplying the target concentration (mol/L) by the volume (L) gives the moles you need, and multiplying by the molecular weight converts moles to grams.

For hydrated salts, use the molecular weight printed on the bottle, which includes the water of hydration (for example, MgCl₂·6H₂O is 203.30 g/mol, not 95.21).

How it works

The amount of solute stays the same when you dilute, so concentration × volume before equals concentration × volume after. Rearranging gives the stock volume. The rest of the final volume is diluent (water, buffer or medium).

Volumes under about 1 µL are hard to pipette accurately. Make an intermediate dilution first if V1 comes out that small.

How it works

X notation is a concentration multiplier: 50X TAE is fifty times stronger than the working 1X buffer, so you dilute it 1 in 50. Percent solutions dilute the same way, using C1V1 = C2V2.

For % w/v, 1% means 1 g per 100 mL, so a 2% agarose or SDS solution needs 2 g per 100 mL. Ethanol and water shrink slightly when mixed, so for exact work bring the mix up to the final volume rather than adding a fixed volume of water.

How it works

A buffer resists pH change best within about one unit of its pKa. The Henderson–Hasselbalch equation gives the ratio of the base form (A⁻) to the acid form (HA) needed for your target pH, which this splits into concentrations and amounts for your volume.

Treat the result as a starting point and always finish by adjusting with a pH meter. pKa shifts with temperature and ionic strength; Tris drops about 0.03 pH units per °C, so set its pH at the temperature you will use it.

How it works

Amounts are given for anhydrous salts unless noted. Dissolve the solids in about 80% of the final volume, adjust pH if the recipe calls for it, then bring to volume. Autoclave or filter-sterilise as your protocol requires.

How it works

Each tube gets the same volume of diluent, then a fixed volume is carried from the previous tube, mixed, and carried on. Choosing the transfer as keep ÷ (factor − 1) leaves every tube with the volume you want after its transfer out. The last tube keeps the extra transfer volume, so remove that amount if every tube must be equal.

Use a fresh tip at each step and mix thoroughly; small carry-over errors compound down the series.

How it works

Each component's volume per reaction is final ÷ stock × reaction volume. Water fills the remainder. Template is added to each tube separately, so it is left out of the master mix total but counted when calculating water.

The extra percentage covers liquid left in tips and tube walls. 10% is typical; use more for viscous mixes or many reactions.

How it works

The nearest-neighbor method adds up the stacking energy of each neighbouring base pair, which predicts Tm far better than counting G/C bases alone. The result depends on salt and primer concentration, so set them to match your buffer. Polymerase vendors often use their own buffer-specific formulas, so expect a few degrees of difference.

A good primer pair usually has 40–60% GC, 18–25 bases, and Tm values within 5 °C of each other. A starting annealing temperature is about 3–5 °C below the lower Tm.

How it works

Subtracting the reference gene Ct normalises for how much cDNA went into each well (ΔCt). Comparing the treated ΔCt with the control ΔCt gives ΔΔCt, and since each cycle doubles the product, the fold change is 2 raised to −ΔΔCt.

This assumes both primer pairs amplify with close to 100% efficiency. Check that with the qPCR efficiency calculator; if efficiencies differ by more than about 10%, use the Pfaffl method instead.

How it works

Ct is plotted against the log of relative template amount and fitted with a straight line. A perfect reaction doubles every cycle, giving a slope of −3.32 for 10-fold dilutions and 100% efficiency. Acceptable assays are usually 90–110% with R² above 0.98.

Efficiency above 110% often means inhibition in the most concentrated points or primer-dimers; try leaving out the top dilution.

How it works

Ligation depends on the number of DNA ends, not their mass. A short insert has more molecules per nanogram than a long vector, so the mass is scaled by the length ratio. 3:1 is a common starting point; very short inserts often work better at 5:1 or higher.

How it works

Gel percentage is w/v, so a 1% gel has 1 g of agarose per 100 mL of buffer. Higher percentages resolve smaller fragments. Use the same buffer (TAE or TBE) in the gel and the tank. A mini gel tray usually takes 30–50 mL.

Suggested percentages are starting points: about 0.7–0.8% for fragments above 5 kb, 1% for 0.5–5 kb, 1.5% for 0.3–1 kb and 2% or more below 500 bp.

How it works

Only part of the transformation goes on the plate, so the DNA on the plate is the DNA transformed × (volume plated ÷ total volume) ÷ dilution. Dividing the colony count by that amount in µg gives efficiency. Test with a small amount (about 10 pg–1 ng) of an intact supercoiled plasmid such as pUC19.

Home-made chemically competent cells are often around 10⁶–10⁷ cfu/µg; commercial high-efficiency cells reach 10⁸–10⁹.

How it works

An A260 of 1.0 (1 cm path) corresponds to about 50 ng/µL of double-stranded DNA, 40 ng/µL of RNA or 33 ng/µL of single-stranded DNA. Readings are most reliable between 0.1 and 1.0.

A260/280 near 1.8 suggests clean DNA and near 2.0 clean RNA. Lower ratios usually point to protein or phenol carryover.

How it works

1 µM equals 1 pmol/µL, so dividing the nanomoles by the target concentration and multiplying by 1000 gives microlitres. A 100 µM stock is the usual choice: concentrated enough to store, easy to dilute to a 10 µM working stock.

Spin the tube before opening so the pellet is at the bottom, and resuspend in low-EDTA TE (pH 8) for long-term storage.

How it works

Sequencers count molecules, so libraries are pooled by molarity, not mass. Concentration in nM is ng/µL × 10⁶ ÷ (660 × average size in bp), using about 660 g/mol per base pair. Each library then contributes an equal share of the pool's molarity, and buffer or water makes up the rest of the volume.

Use the average size from a Bioanalyzer or TapeStation trace including adapters, and a fluorometric concentration (Qubit) rather than A260. For the most accurate loading, confirm the final pool by qPCR.

How it works

An average base pair of double-stranded DNA weighs about 650 g/mol. Dividing the mass by the molecule's molar mass gives moles, and multiplying by Avogadro's number gives the copy number.

How it works

Each large square of a Neubauer chamber holds 0.1 µL (1 mm × 1 mm × 0.1 mm), so the mean count per square × 10,000 gives cells per mL, before correcting for dilution. Trypan blue enters only cells with damaged membranes, so blue cells are counted as dead.

Aim for 20–100 cells per square for a reliable count; dilute the sample further if they overlap.

How it works

The calculator makes one mix for all wells: the total cells needed divided by your counted concentration gives the suspension volume, and medium makes up the rest. Mix well and pipette the per-well volume into each well.

How it works

During exponential growth, cell number doubles at a constant rate. The number of doublings is log₂(N ÷ N₀), and the elapsed time divided by that number gives the doubling time. Counts taken during lag or stationary phase will overestimate it.

How it works

Absorbance at 280 nm comes mainly from tryptophan and tyrosine, so each protein has its own extinction coefficient. Dividing absorbance by ε and the path length gives molar concentration, and multiplying by molecular weight converts it to mg/mL.

Nucleic acid contamination inflates A280; check that A260/A280 is around 0.6 for a pure protein.

How it works

The force on a sample depends on both speed and distance from the centre, so the same RPM gives different g-force on different rotors. Protocols should state × g; this converts it to the speed setting for your rotor's radius.