Core bench skills and instruments: balances, glassware, pipetting, buffers, centrifuges, chromatography, spectrophotometry, calibration, microscopy and aseptic technique.

Lab Techniques & Instrumentation

Draw a 12-place rotor as a clock face with positions 0 to 11.

Mark a balanced pattern for 2, 3, 4 and 6 equal tubes, one drawing each.

Mark a balanced pattern for 5 equal tubes without a balance tube (hint: combine a triangle and an opposite pair that do not overlap).

Mark a balanced pattern for 7 equal tubes (hint: a triangle plus two opposite pairs).

Calculate the RCF for a rotor with r = 8.5 cm at 13,000 rpm using RCF = 1.118e-5 x r x rpm^2.

A method says 500 x g for 5 minutes. Calculate the rpm for a rotor with r = 12 cm (rearrange the formula: rpm = square root of RCF / (1.118e-5 x r)).

Write a five-line pre-run checklist for a benchtop centrifuge in your own words (tubes, balance, lid, settings, stopping).

If you are taking a lab course, ask your instructor to check your balancing diagrams and to show you the centrifuge used in your teaching lab.

> **Awareness level only.** This lesson explains the ideas so you recognise them. It does not qualify you to do this work. Before you use lab equipment or handle chemicals, biological materials or patient samples for real, you need the training your employer or institution requires, and you must work under supervision until they sign you off.


## Why it matters

Centrifuges separate things by density: cells from culture medium, plasma from blood cells, a DNA pellet from ethanol, a precipitate from its solution. They are among the most common instruments in any biology, chemistry or clinical lab, and among the most dangerous when misused. A rotor spinning at thousands of revolutions per minute stores a lot of energy. An unbalanced load can make the centrifuge shake, "walk" across the bench, or in the worst case fail the rotor. Broken tubes can release aerosols of whatever was inside.

## How it works

Spinning a sample creates a force that pushes denser particles outward, toward the bottom of the tube. They collect as a **pellet**. The liquid above is the **supernatant**. Separation depends on particle size and density, the liquid's viscosity, the force and the time.

### RPM versus RCF

Protocols should state the **relative centrifugal force (RCF)** in multiples of gravity (× g), not the speed in revolutions per minute (rpm). The same rpm gives different forces in rotors of different radius:

```text
RCF = 1.118e-5 x r x rpm^2
```

where r is the rotor radius in centimetres (to the middle or bottom of the tube, as your protocol specifies).

**Worked example.** A rotor with r = 10 cm at 3,000 rpm:

```text
RCF = 1.118e-5 x 10 x 3000^2 = 1.118e-5 x 10 x 9,000,000 = about 1,006 x g
```

The same 3,000 rpm in a rotor with r = 15 cm gives about 1,509 × g. If a method says "1,000 × g for 10 minutes", you calculate the rpm for your rotor, or use the instrument's RCF setting.

## Types of centrifuge

| Type | Typical maximum | Common uses |
|---|---|---|
| Microcentrifuge | about 15,000 to 21,000 × g | 1.5 and 2 mL tubes, DNA and protein preps |
| Benchtop centrifuge | a few thousand × g | Blood tubes, cell pellets, 15 and 50 mL tubes |
| High-speed floor centrifuge | tens of thousands × g | Larger volumes, subcellular fractions |
| Ultracentrifuge | hundreds of thousands × g | Viruses, ribosomes, density gradients |

Ultracentrifuges and high-speed rotors have strict rules: rotor logs, speed derating as rotors age, and specific training. Do not use one until you are trained on it.

## Balancing a load

A centrifuge must be balanced: tubes of **equal mass** in positions **symmetric about the centre**.

1. Use matching tubes, filled to matching volumes. For critical runs, check pairs on a balance.
2. Place each tube directly opposite another tube of equal mass.
3. With an odd number of samples, add a **balance tube** filled with water to the same mass, or use a symmetric pattern. In a 6-place rotor, three tubes at every other position (a triangle) are balanced.
4. Swing-bucket rotors: all buckets must be in place, even empty ones, and paired buckets should match.

**Worked example.** You have 5 tubes and a 12-place rotor. Number the positions 0 to 11 like a clock. A triangle of positions 0, 4 and 8 is balanced, and so is the opposite pair 3 and 9. Together, positions 0, 3, 4, 8, 9 hold five equal tubes in balance. Without a pattern like this, add a sixth water tube opposite the odd one.

## Running it safely

- Inspect tubes for cracks and use tubes rated for the speed.
- Close tube caps, and for biological material use **sealed rotors or safety cups (aerosol-tight buckets)**. Open them in a biosafety cabinet if your SOP says so.
- Close the lid, set time, speed or RCF, and temperature, then start.
- Stay nearby until it reaches speed. If you hear unusual noise or feel vibration, stop it.
- **Never open the lid** until the rotor has completely stopped, and never slow it by hand.
- If a tube breaks, follow your lab's spill procedure. For biological material this usually means waiting before opening to let aerosols settle, then disinfecting.
- Log use where required.

## Common mistakes

- Following an rpm value written for someone else's rotor.
- Balancing by tube count instead of mass (a full tube opposite a half-full one).
- Leaving out a swing bucket.
- Opening the lid during the run down.
- Using tubes past their rated speed.

## Quick recap

- Protocols should specify RCF (× g); convert with RCF = 1.118e-5 x r x rpm^2.
- Balance by mass and symmetry; use a water balance tube when needed.
- Sealed rotors or safety cups for biological samples; never open until stopped.
- Higher-speed instruments need specific training and logs.

## Try it

Open the **Do** tab to work out balanced loading patterns and RCF conversions on paper.


Balance and run a centrifuge safely, and convert between rpm and relative centrifugal force (× g).

Centrifugation

How do you use a Centrifuge? A step-by-step guide!

Basic Centrifugation

## Why it matters

Chromatography is how labs separate and measure the parts of a mixture: drugs in a tablet, pesticides in water, amino acids in a protein digest, drugs of abuse in urine. Chemists, biochemists and medical laboratory staff all use it, and many technician jobs are built around running and maintaining HPLC or GC systems. This lesson explains the principles so you can follow what an instrument is doing and read a chromatogram.

## The core idea

Every chromatography method has two phases:

- A **stationary phase** that stays put: a coated plate, a packed column or a coated capillary.
- A **mobile phase** that moves through it: a solvent (liquid chromatography) or a gas (gas chromatography).

Each compound in the sample divides its time between the two phases. Compounds that interact more strongly with the stationary phase move more slowly and come out later. That difference in speed separates them.

## Thin-layer chromatography (TLC)

TLC is the simplest form. The stationary phase is a thin layer of silica on a plate. You spot the sample near the bottom, stand the plate in a shallow pool of solvent in a closed chamber, and let the solvent climb by capillary action. When the solvent front is near the top, you mark it, dry the plate, and see the spots (often under UV light or with a stain).

The **retention factor (Rf)** describes how far a spot moved:

```text
Rf = distance travelled by the spot / distance travelled by the solvent front
```

**Worked example.** The solvent front moved 8.0 cm. Spot A moved 2.0 cm (Rf 0.25) and spot B moved 6.0 cm (Rf 0.75). On normal silica, which is polar, the more polar compound sticks more and has the lower Rf, so A is more polar than B. TLC is quick and cheap: it is used to check whether a reaction is finished or whether a sample is pure. It is mostly qualitative.

## High-performance liquid chromatography (HPLC)

HPLC pumps liquid mobile phase at high pressure through a column packed with very small particles. The parts of a system are:

1. **Solvent reservoirs and pump:** deliver a steady flow, either at a fixed composition (isocratic) or a changing one (gradient).
2. **Autosampler:** injects a precise volume of each sample.
3. **Column:** the stationary phase. The most common is **reversed-phase C18**: a non-polar coating, with a polar mobile phase such as water and acetonitrile or methanol. Non-polar compounds are retained longer.
4. **Detector:** often UV-visible absorbance, sometimes fluorescence or a mass spectrometer (LC-MS).
5. **Data system:** draws the chromatogram.

## Gas chromatography (GC)

GC uses an inert **carrier gas** (helium, hydrogen or nitrogen) as the mobile phase and a long, thin coated capillary column in a temperature-controlled oven. The sample must be **volatile** (or made volatile) and stable when heated. Compounds separate by boiling point and by interaction with the coating. Common detectors are the **flame ionisation detector (FID)** for organic compounds and the **mass spectrometer (GC-MS)**, which also identifies compounds by their fragmentation pattern. GC suits solvents, fuels, fragrances, blood alcohol and many environmental contaminants.

## Reading a chromatogram

A chromatogram plots detector signal against time. Each peak is a compound (or several compounds that did not separate).

- **Retention time:** when the peak appears. Under fixed conditions it helps identify the compound, by comparison with a standard.
- **Peak area (or height):** proportional to the amount. With a calibration curve from standards, area gives concentration.
- **Resolution:** how well two neighbouring peaks are separated. Overlapping peaks give unreliable areas.
- **Peak shape:** tailing or split peaks often point to column or method problems.

Regulated labs run a **system suitability test** before samples: a standard injection that must meet limits for retention time, resolution, peak shape and repeatability. If it fails, you do not run samples.

## Choosing a method

| Question | Usually points to |
|---|---|
| Quick check of reaction progress or purity | TLC |
| Non-volatile or heat-sensitive compounds (most drugs, proteins, sugars) | HPLC |
| Volatile compounds (solvents, fuels, alcohols) | GC |
| Need to identify unknowns with high confidence | LC-MS or GC-MS |

## Common mistakes

- Treating retention time alone as proof of identity. Two compounds can share a retention time; MS or a second method confirms it.
- Running samples after a failed system suitability test.
- Letting the solvent pool rise above the TLC spots (the sample dissolves into the pool).
- Integrating peaks inconsistently, which changes the reported amount.

## Quick recap

- Separation happens because compounds split their time differently between a stationary and a mobile phase.
- TLC gives Rf; HPLC uses liquid under pressure, often on reversed-phase C18; GC uses a carrier gas for volatile compounds.
- Retention time helps identify, peak area quantifies, and resolution tells you whether you can trust the areas.
- System suitability must pass before samples run.

Hands-on chromatography is learned in your teaching lab and on the job, under supervision.

## Try it

This lesson is quiz-only. Open the **Do** tab to check your understanding.


Explain how TLC, HPLC and GC separate mixtures, and read retention time, peak area and resolution on a chromatogram.

Chromatography basics

Thin Layer Chromatography (TLC)

Quickly Understand High Performance Liquid Chromatography (HPLC)

> **Awareness level only.** This lesson explains the ideas so you recognise them. It does not qualify you to do this work. Before you use lab equipment or handle chemicals, biological materials or patient samples for real, you need the training your employer or institution requires, and you must work under supervision until they sign you off.


## Why it matters

Spectrophotometry is one of the most used measurements in science. Labs use it to measure protein and DNA concentration, follow enzyme reactions, run colour-based clinical tests and check water quality. The instrument is simple to operate. Getting a trustworthy number depends on understanding the law behind it and building a good **standard curve**.

## Absorbance and transmittance

A spectrophotometer shines light of one wavelength through a sample in a **cuvette** and measures how much gets through.

- **Transmittance (T)** is the fraction of light that passes through: T = I / I0.
- **Absorbance (A)** is defined as A = -log10(T).

| Transmittance | Absorbance |
|---|---|
| 100 percent (1.0) | 0 |
| 10 percent (0.1) | 1 |
| 1 percent (0.01) | 2 |

Absorbance is used because it is proportional to concentration.

## The Beer-Lambert law

```text
A = epsilon x b x c
```

- **epsilon** (molar absorptivity) is how strongly the compound absorbs at that wavelength, in L/(mol·cm).
- **b** is the path length, usually 1 cm for a standard cuvette.
- **c** is the concentration in mol/L.

**Worked example.** NADH absorbs at 340 nm with epsilon of about 6,220 L/(mol·cm). A reading of A = 0.311 in a 1 cm cuvette gives c = 0.311 / (6,220 x 1) = 5.0 x 10^-5 mol/L, or 50 µmol/L.

## Choosing a wavelength and a blank

- Measure at the compound's **absorbance maximum (lambda max)**, where sensitivity is highest and small wavelength errors matter least.
- **Blank** the instrument with a cuvette of everything except the analyte (the solvent or reagent blank). This sets A = 0 and removes the absorbance of the solvent, the reagents and the cuvette.

## The standard curve

Epsilon is not always known, and real assays include reagents that change it. Labs therefore build a **standard curve** (calibration curve):

1. Prepare standards of known concentration that **bracket** the expected unknowns.
2. Measure the absorbance of each, after blanking.
3. Fit a straight line: A = slope x c + intercept.
4. Measure the unknown and solve for concentration: c = (A - intercept) / slope.

**Worked example.** Standards of 0, 2, 4, 6, 8 and 10 mg/L read 0.002, 0.101, 0.199, 0.304, 0.398 and 0.502. A least-squares fit gives a slope of about 0.0499 per mg/L and an intercept of about 0.0013. An unknown reading 0.250 is (0.250 - 0.0013) / 0.0499 = about 4.98 mg/L.

If you diluted the unknown 1 in 5 before reading it, multiply by the **dilution factor**: 4.98 x 5 = 24.9 mg/L in the original sample.

### Stay in the linear range

Beer-Lambert holds only over a range. At high absorbance (often above about 1 to 1.5, depending on the instrument) very little light reaches the detector, stray light dominates and the curve bends. If an unknown reads above your highest standard, **dilute it and re-read**; do not extrapolate the line.

## Cuvette handling

- Hold cuvettes by the frosted or ribbed sides; fingerprints on the clear faces absorb light.
- Use quartz cuvettes below about 340 nm (UV); plastic and glass absorb UV light.
- Wipe the outside with lint-free tissue, check for bubbles and insert in the same orientation every time.

## Common mistakes

- Forgetting to blank, or blanking with the wrong solution.
- Extrapolating beyond the highest standard.
- Forgetting the dilution factor.
- Reading at the wrong wavelength.
- Using plastic cuvettes for UV work.

## Quick recap

- A = -log10(T), and A = epsilon x b x c.
- Blank first, measure at lambda max, keep readings in the linear range.
- Build a standard curve that brackets the unknowns; c = (A - intercept) / slope, then multiply by any dilution factor.

## Try it

Open the **Do** tab and finish a Python standard curve. The fit is done for you with `statistics.linear_regression`; you write the line that turns an absorbance into a concentration.


Use the Beer-Lambert law and a standard curve to turn absorbance readings into concentrations.

Spectrophotometry

Spectrophotometry and Beer's Law

Spectrophotometry and the Beer–Lambert Law | AP Chemistry | Khan Academy

Balance and run a centrifuge, understand TLC, HPLC and GC, and build a Beer-Lambert standard curve.