An intermediate course for future biochemists, biological technicians, medical laboratory technologists, chemists and materials scientists. The first module covers core bench skills: weighing and measuring to the stated precision, using a micropipette accurately, and calculating and preparing solutions and buffers. The second module explains separation and analysis: balancing and running a centrifuge, how TLC, HPLC and GC separate mixtures, and building a Beer-Lambert standard curve for spectrophotometry. The third module covers instruments and upkeep: calibrating a pH meter and keeping the log, comparing fluorescence and electron microscopy, and the habits of aseptic technique. Python activities are fill-in-the-blank, so no programming background is needed. Everything here is awareness level: it does not replace supervised training in your teaching lab or workplace, and every activity is a paper, data or plain-water exercise that is safe to do at home. Lab careers also have formal requirements, such as a NAACLS-accredited program and ASCP certification for medical laboratory scientists, a chemistry or biology degree with lab coursework, and on-the-job GMP training in regulated manufacturing.

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

Lab Techniques & Instrumentation

Gather a kitchen scale that reads to 1 g (or 0.1 g), a clear measuring jug or cup with millilitre markings, a small bowl, a notebook and a pen.

Put the bowl on the scale and press tare (zero). Write down what tare does in your own words.

Fill the jug to the 100 mL mark with tap water. Crouch so your eye is level with the mark and read the bottom of the curved surface. Then look from above and note how the reading seems to change (parallax).

Pour the 100 mL into the tared bowl and record the mass in grams. Water is close to 1.00 g per mL at room temperature, so 100 mL should weigh about 100 g.

Empty and dry the bowl, then repeat the measurement four more times. Record all five masses in a table, writing down every digit the scale shows.

Calculate the mean of your five masses and the difference between the mean and 100 g. That difference is your accuracy (bias).

Calculate the range (largest minus smallest). That is a simple measure of your precision.

Write one sentence on which source of error you think was biggest: the jug's markings, your reading of the meniscus, or the scale's readability.

Optional, only in your teaching lab and only if your instructor sets it up: repeat the exercise with a Class A volumetric pipette and an analytical balance, and compare your precision.

> **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

Almost every lab result starts with a mass or a volume. If the 250.0 mg you weighed was really 247 mg, or the 100 mL flask was filled past the line, every number downstream carries that error: the concentration of your standard, the calibration curve built from it, and the patient or product result read from that curve. Balances and volumetric glassware are the first instruments you will be trained on, and the habits you build with them follow you through your whole career.

## Accuracy, precision and readability

- **Accuracy** is how close a measurement is to the true value.
- **Precision** is how close repeated measurements are to each other.
- **Readability** is the smallest step the display shows.

A balance can be precise but inaccurate (it always reads 0.3 mg high) or have fine readability but poor precision (the last digit jumps around in a draught). Calibration and good technique address different parts of this.

## Balances

| Type | Typical readability | Typical use |
|---|---|---|
| Top-loading (precision) balance | 0.01 g or 0.001 g | Buffer salts, media, larger masses |
| Analytical balance | 0.0001 g (0.1 mg) | Standards, reference materials |
| Microbalance | 0.001 mg or finer | Very small samples, filters |

Good weighing practice on an analytical balance:

1. **Check it is level.** Most balances have a bubble level. An unlevel balance reads wrong.
2. **Check the daily verification.** Many labs weigh a certified check weight each day and log it. If the log shows a failure, the balance is out of service.
3. **Close the draught shield doors** before reading. Air currents move the last digit.
4. **Let warm or cold items reach room temperature.** A warm beaker creates convection currents and reads light.
5. **Use a weighing boat or paper, and tare it.** Taring sets the display to zero with the container on the pan.
6. **Weigh by difference** for hygroscopic or valuable material: weigh the closed vial, remove some material, weigh the vial again. The mass transferred is the difference, and nothing left on a weighing boat is lost from the calculation.
7. **Keep the pan clean.** Spilled material changes later readings and can damage the mechanism.

### Worked example: weighing by difference

| Reading | Mass (g) |
|---|---|
| Vial with sample, before | 12.4831 |
| Vial with sample, after transfer | 12.2337 |
| **Transferred** | **0.2494** |

You report 0.2494 g (249.4 mg), with all four decimal places the balance gave you.

## Volumetric glassware

Glassware falls into two groups:

- **Volumetric (precise):** volumetric flasks, volumetric (transfer) pipettes and burettes. They are calibrated to a single volume or a fine scale.
- **Approximate:** beakers, Erlenmeyer flasks and, to a lesser extent, graduated cylinders. Beaker markings are only rough guides, often within about 5 percent. Never use a beaker to measure a volume that matters.

Volumetric glassware is made to a **class**. Class A has the tightest tolerances, and Class B tolerances are roughly twice as wide. The tolerance is printed on the glass. For example, a 100 mL Class A volumetric flask is typically marked ±0.08 mL.

Two markings tell you how the glass was calibrated:

- **TC (to contain):** the flask holds the stated volume when filled to the line. Volumetric flasks are TC.
- **TD (to deliver):** the stated volume comes out when the pipette drains, leaving a small film behind. Most transfer pipettes are TD, and you do not blow out the last drop unless the pipette is marked for it.

### Reading a meniscus

Water forms a curved surface (a meniscus) that dips in the middle. Read the **bottom of the meniscus** with your eye level with the line. Looking from above or below causes **parallax error**. A white card with a dark stripe held behind the glass makes the bottom of the curve easier to see.

### Significant figures

Record every certain digit plus one estimated digit. A 50 mL burette has 0.1 mL graduations, so you read it to 0.01 mL, for example 23.46 mL. Writing "23.5 mL" throws away information you actually measured.

## Common mistakes

- Measuring an important volume with a beaker or graduated cylinder instead of a volumetric pipette or flask.
- Filling a volumetric flask to the line before the solid has fully dissolved, then topping up later.
- Reading the meniscus from above.
- Weighing a sample straight from the oven.
- Rounding a balance reading in the notebook.
- Using a balance that failed its daily check.

## Beyond this course

This course teaches the knowledge behind bench work. Real bench skills come from supervised practice, and the careers it serves have formal requirements:

- **Medical laboratory technologist / scientist:** a NAACLS-accredited program with clinical rotations (https://www.naacls.org), ASCP Board of Certification MLS certification (https://www.ascp.org/content/board-of-certification), and a state licence in states that require one.
- **Biological technician:** an associate or bachelor's degree in biology or biotechnology with supervised lab hours. In regulated manufacturing, GMP training happens on the job (FDA CGMP: https://www.fda.gov/drugs/pharmaceutical-quality-resources/current-good-manufacturing-practice-cgmp-regulations).
- **Chemist:** a bachelor's in chemistry with hands-on lab courses; ACS-approved programs are preferred by many employers (https://www.acs.org/education/policies/acs-approval-program.html).
- **Biochemist or materials scientist:** a bachelor's with real lab or instrument coursework for technician roles; an MS or PhD for independent research.

Requirements vary by state and employer, so check the official sources. (Checked on 2026-10-23.)

## Quick recap

- Accuracy, precision and readability are different properties.
- Level, verify, close the doors, tare, and weigh by difference when it matters.
- Use Class A volumetric glassware for exact volumes, and read the bottom of the meniscus at eye level.
- Record every digit the instrument gives you, plus one estimated digit for scales you read by eye.

Real lab training covers your institution's balances and glassware in person. For lab safety rules, see OSHA's laboratory page, https://www.osha.gov/laboratories (Checked on 2026-10-23.)

## Try it

Open the **Do** tab to practise accuracy, precision and meniscus reading with a kitchen scale, a measuring jug and plain tap water.


Weigh and measure to the stated precision: balances, weighing by difference, volumetric glassware and reading a meniscus.

Balances and volumetric glassware

How to Use an Analytical Balance

The Volumetric Pipet and Pipetting Technique

Copy these ten weighings for a P200 set to 50 µL into a table or spreadsheet: 49.2, 50.4, 49.8, 50.1, 48.9, 50.3, 49.6, 50.0, 49.4, 50.2 (mg).

Calculate the mean mass. Convert it to volume using 1.003 µL per mg.

Calculate the systematic error in µL and as a percentage of 50 µL.

Calculate the sample standard deviation and the CV (standard deviation divided by mean, as a percentage).

Suppose your lab's limits for this pipette at 50 µL are ±1.0 percent systematic error and 0.6 percent CV. Decide whether the pipette passes, and write one sentence explaining why.

List three technique errors from the lesson that could explain a high CV.

In your teaching lab, with your instructor's permission and supervision, practise forward pipetting plain water: set a volume, attach a tip, find the first and second stops, aspirate vertically and dispense against a tube wall ten times.

> **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

Biology and clinical labs move small volumes all day: 2 µL of DNA into a reaction, 50 µL of serum into a cuvette, 200 µL of standard into each well of a plate. A micropipette makes this possible, but it is easy to use badly. Holding it at an angle, plunging too fast or using the wrong stop can each cause errors of several percent, and those errors are invisible until a control fails.

## How an air-displacement micropipette works

Most lab micropipettes are **air-displacement** pipettes. The plunger moves a piston inside the barrel. When you release the plunger, the piston rises and the air cushion beneath it pulls liquid into a disposable tip. The liquid never touches the pipette itself, only the tip.

Pipettes come in sizes. Common names follow the maximum volume:

| Pipette | Typical range |
|---|---|
| P2 | 0.2 to 2 µL |
| P10 | 1 to 10 µL |
| P20 | 2 to 20 µL |
| P200 | 20 to 200 µL |
| P1000 | 100 to 1000 µL |

Choose the **smallest pipette that covers your volume**. Accuracy and precision are best in the upper part of a pipette's range. Pipetting 15 µL with a P20 is better than with a P200. Never turn the volume dial past the pipette's limits: it can damage the mechanism.

## The two stops

Press the plunger slowly and you will feel two points of resistance:

- **First stop:** the calibrated volume. You press to here before aspirating.
- **Second stop (blow-out):** pushes out the last drop when dispensing.

### Forward pipetting (the standard method)

1. Set the volume and attach a fresh tip firmly.
2. Hold the pipette vertically. Press the plunger to the **first stop**.
3. Immerse the tip just below the surface, a few millimetres (about 1 mm for very small volumes, a few mm for large ones). Too deep coats the outside of the tip with extra liquid.
4. Release the plunger **slowly and smoothly**. Wait a second for the liquid to finish entering the tip.
5. Withdraw the tip, touching it against the wall of the vessel if your SOP allows, to remove drops on the outside.
6. Dispense by pressing to the first stop with the tip against the wall of the receiving vessel, then to the **second stop** to blow out the rest.
7. Keep the plunger pressed while you remove the tip from the liquid, then release it. Eject the tip.

### Reverse pipetting

For viscous liquids (glycerol, serum) or liquids that foam (detergent solutions), labs often use **reverse pipetting**: press to the second stop before aspirating, then dispense only to the first stop, leaving the excess in the tip. It reduces bubbles and film errors. Your SOP will say which method to use.

### Habits that improve accuracy

- **Pre-wet the tip** by aspirating and dispensing the liquid two or three times before the measured transfer.
- Let the pipette, tips and liquid reach the same temperature.
- Keep a consistent rhythm and plunger speed.
- Change tips between samples to avoid carry-over.
- Never lay a pipette on its side or turn it upside down with liquid in the tip: liquid can run into the barrel and contaminate it.

## Checking a pipette: the gravimetric method

Labs check pipettes by weighing water, because 1 µL of water at about 20 °C weighs very close to 1 mg. The full procedure (ISO 8655) corrects for temperature and air pressure, but the idea is simple.

### Worked example

Ten 100 µL dispenses of water weigh (in mg): 99.6, 100.1, 99.8, 99.9, 100.2, 99.7, 99.9, 100.0, 99.8, 99.6.

- **Mean:** 99.86 mg. Converting with a factor of about 1.003 µL per mg gives about 100.16 µL.
- **Systematic error (accuracy):** 100.16 minus 100 = +0.16 µL, or +0.16 percent.
- **Random error (precision):** the standard deviation is about 0.20 mg, so the coefficient of variation (CV) is 0.20 / 99.86 = about 0.20 percent.

Your lab compares these numbers with acceptance limits for that pipette size. If a pipette fails, it is labelled out of service and sent for adjustment.

## Common mistakes

- Aspirating at an angle instead of vertically.
- Pressing past the first stop before aspirating (this draws up too much in forward pipetting).
- Letting the plunger snap back, which splashes liquid into the barrel and forms bubbles.
- Using a P1000 at 50 µL.
- Reusing a tip between different samples.
- Skipping the scheduled pipette check because the pipette "seems fine".

## Quick recap

- Pick the smallest pipette that covers the volume, and stay within its range.
- First stop to aspirate, second stop to blow out (forward pipetting).
- Vertical, shallow, slow and smooth; pre-wet the tip; change tips.
- Pipettes are checked by weighing water: mean gives accuracy, CV gives precision.

## Try it

Open the **Do** tab to analyse a pipette check on paper, then practise the plunger motions with plain water if you have access to a teaching lab.


Use a micropipette accurately: choosing the right size, the two stops, forward and reverse pipetting, and gravimetric checks.

Pipetting

How to Use a Micropipette

Proper Pipetting Technique

> **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

Buffers and reagent solutions are the background of nearly every assay. An enzyme assay at the wrong pH, a cell wash at the wrong salt concentration or a standard made at the wrong molarity can make a whole run invalid. The good news is that the maths is short and the same few formulas cover most lab work.

## Molarity and the core formula

**Molarity (M)** is moles of solute per litre of solution. To find the mass you need:

```text
mass (g) = molarity (mol/L) x volume (L) x molar mass (g/mol)
```

### Worked example: 500 mL of 0.150 M NaCl

NaCl has a molar mass of 58.44 g/mol.

```text
mass = 0.150 x 0.500 x 58.44 = 4.383 g
```

Weigh 4.383 g, dissolve it in less than 500 mL of water in a beaker, transfer quantitatively to a 500 mL volumetric flask (rinsing the beaker into the flask), and make up to the line.

**Check the label for the exact form.** Many salts come as hydrates. Magnesium chloride hexahydrate (MgCl2·6H2O, 203.30 g/mol) needs more than twice the mass of anhydrous MgCl2 (95.21 g/mol) for the same molarity. Using the wrong molar mass is one of the most common prep errors.

## Dilutions

When you dilute a stock, the amount of solute stays the same:

```text
C1 x V1 = C2 x V2
```

To make 250 mL of 0.10 M from a 2.0 M stock: V1 = 0.10 x 250 / 2.0 = 12.5 mL of stock, made up to 250 mL. A **serial dilution** repeats a fixed step (for example 1:10 each time) to reach very low concentrations accurately.

Other units you will meet: **percent w/v** (grams per 100 mL, so 0.9 percent NaCl is 0.9 g per 100 mL) and **mg/mL** or **µg/mL** for standards.

## Buffers

A **buffer** resists changes in pH. It is a mixture of a weak acid (HA) and its conjugate base (A-). The **Henderson-Hasselbalch equation** links pH to their ratio:

```text
pH = pKa + log10( [A-] / [HA] )
```

Rearranged, the base-to-acid ratio you need is:

```text
[A-] / [HA] = 10 ** (pH - pKa)
```

A buffer works best within about **one pH unit of its pKa**. Outside that range it has little capacity to absorb added acid or base.

| Buffer system | pKa (about 25 °C) | Useful range |
|---|---|---|
| Acetate | 4.76 | 3.8 to 5.8 |
| Phosphate (H2PO4- / HPO4 2-) | 7.21 | 6.2 to 8.2 |
| Tris | 8.06 | 7.1 to 9.1 |

### Worked example: 1 L of 0.10 M phosphate buffer, pH 7.40

1. Ratio: 10 ** (7.40 - 7.21) = 10 ** 0.19 = 1.549.
2. Fraction that is base: 1.549 / (1 + 1.549) = 0.608. Base = 0.0608 M, acid = 0.0392 M.
3. Masses for 1 L, using anhydrous salts:
   - Na2HPO4 (base, 141.96 g/mol): 0.0608 x 1 x 141.96 = 8.63 g
   - NaH2PO4 (acid, 119.98 g/mol): 0.0392 x 1 x 119.98 = about 4.71 g (4.707 g before rounding the molarity)

The calculation gets you close. Real solutions differ from the ideal equation because of ionic strength and temperature, so the standard practice is:

- Dissolve the components in about 80 percent of the final volume.
- Measure with a **calibrated pH meter** at the temperature you will use the buffer, and adjust with small additions of acid or base as your SOP specifies.
- Make up to the final volume **after** adjusting the pH.
- Label it: name, concentration, pH, date, your initials, expiry and lot numbers of the components.

Tris is a classic trap: its pKa falls by about 0.03 units per °C, so a Tris buffer set to pH 8.0 at room temperature is noticeably more alkaline in a cold room.

## Common mistakes

- Using the molar mass of the anhydrous salt when the bottle is a hydrate (or the reverse).
- Making up to volume before adjusting the pH.
- Choosing a buffer whose pKa is more than one unit away from the target pH.
- Setting the pH at room temperature for a buffer used at 4 °C or 37 °C.
- Leaving bottles unlabelled "just for today".

## Quick recap

- mass = M x V x molar mass; dilute with C1V1 = C2V2.
- Henderson-Hasselbalch gives the base-to-acid ratio: 10 ** (pH - pKa).
- Pick a buffer with a pKa within one unit of your target, then confirm with a calibrated pH meter.
- Label every solution completely.

## Try it

Open the **Do** tab and finish a small Python buffer calculator. The code already runs: you write one line, the Henderson-Hasselbalch ratio.


Calculate masses, dilutions and Henderson-Hasselbalch ratios to prepare solutions and buffers.

Preparing solutions and buffers

Solution Preparation

Preparing Tris Buffer

Weigh and measure to the stated precision, use a micropipette accurately, and calculate and prepare solutions and buffers.

Core Bench Skills

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.

Separation and Analysis

Set up a calibration log with these columns: instrument ID, date and time, standards with lot and expiry, temperature, slope (percent), offset (mV), limits, pass or fail, action, initials.

Calibration 1 at 25 °C: pH 7.00 reads +4 mV and pH 4.01 reads +175 mV. Calculate the slope in mV per pH unit and as a percentage of 59.16. Record it in the log.

Calibration 2 at 25 °C: pH 7.00 reads -6 mV and pH 4.01 reads +150 mV. Calculate the slope percentage and record it.

Using limits of 95 to 105 percent slope and ±30 mV offset, mark each calibration pass or fail.

For the failing calibration, write the action you would take: what label goes on the meter, who you tell, and what you check next (cleaning, reconditioning, buffer quality).

Write one sentence on which sample results might need review if the failing meter had been used the previous afternoon.

Find the calibration or service sticker on any instrument you have access to (a thermostat, a scale or, in your teaching lab, a balance or pH meter) and note what information it shows.

If you are in a lab course, ask your instructor to demonstrate a two-point pH calibration and compare their log entry with yours.

> **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

An instrument is only as trustworthy as its last calibration. A pH meter that drifted 0.2 units overnight, a balance that was knocked out of level, or a thermometer that reads 1 °C low can all produce results that look perfectly normal and are wrong. Calibration, verification and maintenance, and the records that prove they happened, are what let a lab stand behind its numbers. Auditors check these logs early in any inspection.

## Three words that are often mixed up

- **Calibration:** comparing an instrument's readings with known reference standards, and usually adjusting it so it reads correctly. Example: calibrating a pH meter with pH 4, 7 and 10 buffers.
- **Verification (or check):** confirming that the instrument still reads within limits, without adjusting it. Example: weighing a certified check weight on a balance each morning.
- **Maintenance:** keeping the instrument working: cleaning, replacing parts, servicing. Some is daily (wiping a balance pan), some yearly by an engineer (preventive maintenance).

**Traceability** means each reference standard is linked, through an unbroken chain of calibrations, to national or international standards (in the US, often through NIST). Certificates for buffers, weights and thermometers state this traceability.

## Calibrating a pH meter

A pH electrode produces a voltage that changes with pH. At 25 °C an ideal electrode changes by **59.16 mV per pH unit**. Real electrodes age, so the meter is calibrated against buffers of known pH.

Typical steps (your SOP and the meter's manual come first):

1. Check the buffers are in date and at room temperature, and note their lot numbers.
2. Rinse the electrode with deionised water and blot (do not wipe) it dry.
3. Place it in **pH 7.00** buffer first (this sets the offset, or zero point). Wait for a stable reading and accept.
4. Rinse, blot, and place it in **pH 4.01** (or 10.01) buffer. This sets the **slope**. For samples across a wide range, use a three-point calibration with 4, 7 and 10.
5. Read the slope and offset the meter reports and compare them with your acceptance limits. A common rule of thumb is a slope of **95 to 105 percent** and an offset within about ±30 mV, but follow your lab's limits.
6. Verify with a buffer not used in the calibration, if your SOP requires it.
7. Store the electrode in its **storage solution** (often KCl), never in deionised water, which damages it over time.
8. **Log it.**

### Worked example: is this electrode acceptable?

At 25 °C the meter reads -2 mV in pH 7.00 buffer and +168 mV in pH 4.01 buffer.

```text
slope = (168 - (-2)) / (7.00 - 4.01) = 170 / 2.99 = 56.9 mV per pH unit
percent of ideal = 56.9 / 59.16 x 100 = 96.1 percent
offset = -2 mV
```

The slope is within 95 to 105 percent and the offset is small, so the calibration passes. A slope of 88 percent would mean the electrode needs cleaning, reconditioning or replacement.

## What goes in the log

A calibration record should let anyone reconstruct what happened. Typical fields:

| Field | Example |
|---|---|
| Instrument ID | PH-03 |
| Date and time | 2026-10-23 08:12 |
| Standards used, lot and expiry | pH 7.00 lot 51A, exp 2027-03; pH 4.01 lot 49C, exp 2027-01 |
| Temperature | 22.4 °C |
| Results | slope 96.1 percent, offset -2 mV |
| Pass or fail against limits | Pass (limits 95 to 105 percent, ±30 mV) |
| Action taken | None |
| Initials | AB |

## When an instrument fails

- Do not use it. Label it **out of service** (a tag or sticker) so others do not either.
- Record the failure and what you did.
- Tell your supervisor.
- Ask the important question: **what was measured since the last good calibration?** Those results may need review or repeat testing. This is often handled as a deviation, which you will meet in the Lab Data & Quality course.

## Preventive maintenance and service

Larger instruments (HPLC, centrifuges, biosafety cabinets, spectrophotometers) have scheduled **preventive maintenance**, often by the manufacturer's engineer, with a sticker showing the next due date. Biosafety cabinets and fume hoods are certified on a schedule. Check the sticker before you use any instrument; an overdue one may not be used for reportable work.

## Common mistakes

- Calibrating with expired or contaminated buffers.
- Wiping a pH electrode (static charge and scratches) instead of blotting it.
- Storing the electrode in deionised water.
- Writing the log at the end of the day from memory.
- Using an instrument with an expired calibration sticker "just this once".

## Quick recap

- Calibration adjusts against traceable standards; verification checks without adjusting; maintenance keeps it working.
- pH: offset at pH 7, slope with 4 or 10; accept a slope around 95 to 105 percent (per your SOP).
- Log every calibration completely and at the time you do it.
- A failed instrument is tagged out, and results since the last good calibration are reviewed.

For traceability and calibration services, see NIST: https://www.nist.gov/calibrations (Checked on 2026-10-23.)

## Try it

Open the **Do** tab to judge calibration data and design a calibration log on paper.


Calibrate a pH meter, judge its slope and offset, and keep a calibration and maintenance log.

Calibration and maintenance

Using a pH Meter

Instrument's Calibration in the Clinical Laboratory: Impact on patient care

## Why it matters

Ordinary light microscopes show cells and tissues, but many lab questions need more. Where in the cell is this protein? Is this bacterium alive? What does a virus or a crack in a metal grain boundary look like at 10 nanometres? Fluorescence and electron microscopy answer those questions, and they are central to biochemistry, cell biology, clinical microbiology and materials science. You may operate these instruments as a technician or depend on core facilities that do.

## The resolution limit of light

**Magnification** makes an image bigger. **Resolution** is the smallest distance at which two points still look separate. Magnifying beyond the resolution limit only makes a blur bigger.

For a light microscope, the Abbe limit is approximately:

```text
d = wavelength / (2 x NA)
```

where NA is the numerical aperture of the objective. With green light (about 500 nm) and a good oil-immersion objective (NA 1.4), d is about 500 / 2.8 = **about 180 nm**. Most viruses and all individual proteins are smaller than that.

## Fluorescence microscopy

A **fluorophore** absorbs light of one wavelength and emits light of a longer wavelength. The difference is the **Stokes shift**. A fluorescence microscope uses this:

1. A bright light source (LED, mercury or xenon lamp, or a laser) passes through an **excitation filter** that selects the wavelength the fluorophore absorbs.
2. A **dichroic mirror** reflects that light onto the sample.
3. The fluorophore emits longer-wavelength light, which passes through the dichroic mirror and an **emission filter** to the camera or eyepiece.
4. Only the labelled structures glow against a dark background.

The excitation filter, dichroic and emission filter are often mounted together as a **filter cube** matched to a fluorophore.

Common labels:

- **DAPI** and **Hoechst** bind DNA and stain nuclei blue.
- **Green fluorescent protein (GFP)** can be genetically fused to a protein so living cells show where it goes.
- **Antibodies tagged with fluorophores** (immunofluorescence) label specific proteins in fixed cells or tissue.

### Confocal and beyond

A widefield fluorescence image includes out-of-focus glow from above and below the focal plane. A **confocal microscope** scans a laser point and uses a **pinhole** to block out-of-focus light, giving sharp **optical sections** that can be stacked into 3D images. **Super-resolution** methods (such as STED, PALM and STORM) use clever tricks with fluorophores to get below the Abbe limit, to tens of nanometres.

### Limits

- **Photobleaching:** fluorophores fade with exposure. Keep light exposure short and image consistently.
- **Phototoxicity:** strong light damages living cells.
- **Autofluorescence:** some tissues glow on their own; controls without the label reveal it.
- You only see what you labelled.

## Electron microscopy

Electrons have a far shorter wavelength than visible light, so electron microscopes resolve down to the nanometre scale or below. The beam must travel in a **vacuum**, so samples must be dry or frozen and stable in vacuum.

| | Transmission EM (TEM) | Scanning EM (SEM) |
|---|---|---|
| How it works | Electrons pass **through** a very thin section | A focused beam scans the **surface**; detectors collect emitted electrons |
| Image | 2D internal structure (organelles, viruses, crystal defects) | 3D-looking surface topography |
| Sample preparation | Fix, embed, cut ultrathin sections (tens of nm), stain with heavy metals | Fix, dry, coat non-conductive samples with a thin metal layer (sputter coating) |
| Typical resolution | Down to below 1 nm | About 1 to 10 nm |

SEM instruments often add **energy-dispersive X-ray spectroscopy (EDS)**, which identifies elements in a spot on the sample. That makes SEM a workhorse in materials science for fracture surfaces, coatings and particles.

**Cryo-electron microscopy (cryo-EM)** freezes samples so fast that water forms glass-like ice instead of crystals, preserving molecules close to their natural state. It is now a major method for determining protein structures.

## Choosing a method

| Question | Better choice |
|---|---|
| Where is protein X in living cells, over time? | Fluorescence (GFP), confocal for 3D |
| What is the internal fine structure of a mitochondrion or virus? | TEM |
| What does the fracture surface of a steel part look like, and what elements are present? | SEM with EDS |
| What is the 3D structure of a purified protein complex? | Cryo-EM |

## Common mistakes

- Confusing magnification with resolution.
- Interpreting a fluorescence image without no-label or no-primary-antibody controls.
- Comparing images taken with different exposure settings.
- Forgetting that EM sample preparation (drying, staining, coating) can create artefacts.

## Quick recap

- Light microscopes are limited to about 200 nm resolution.
- Fluorescence uses excitation and emission filters with labels such as DAPI, GFP and antibodies; confocal adds sharp optical sections.
- TEM looks through thin sections; SEM scans surfaces; both need vacuum and careful preparation.
- Pick the method by the question: living or fixed, inside or surface, molecules or materials.

These instruments are operated only after training by the facility that runs them.

## Try it

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


Compare fluorescence, confocal and electron microscopy, and choose the right method for a question.

Advanced microscopy

Widefield and Confocal Fluorescence Microscopy

Electron Microscopy (TEM and SEM)

Clear a table and wipe it with an ordinary household cleaner. Mark out a "work zone" with tape, with a clean area on one side and a "waste" area on the other.

Set out two clean bottles or jars with lids (the "medium bottle" and the "flask"), a measuring spoon or syringe without a needle, and a label for the flask with today's date and your initials.

Fill the medium bottle with tap water and close it. Write the planned steps on a card before you start.

Wash your hands. Open the medium bottle and hold its lid in your other hand, inner side facing down, without touching the inside.

Transfer a measured amount of water to the flask without touching the rims, then close both containers. Notice whenever your hand passes over an open container.

Put the used spoon or syringe straight into the waste area.

Repeat the whole transfer three times, aiming for no hand-over-open-container moves and the shortest open time.

Write down every moment that would have been a contamination risk in a real lab, and how you would change your layout to avoid it.

If you are taking a lab course, ask your instructor to watch your technique during your next supervised aseptic transfer.

> **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

Aseptic technique is the set of habits that keeps unwanted microbes out of your work, and keeps your work's microbes where they belong. A single contaminating bacterium or fungal spore can ruin a cell culture that took weeks to grow, give a false positive in a clinical culture, or contaminate a batch in a biomanufacturing facility. Good technique also protects you and others. Every biological technician, medical laboratory technologist and biochemist is expected to perform it automatically.

## Sterile versus aseptic

- **Sterile** means free of all living microorganisms. Items are made sterile by autoclaving (steam under pressure), filtration, irradiation or similar.
- **Aseptic** technique means working in a way that keeps sterile items sterile and prevents contamination during handling.

You cannot make your bench sterile, but you can avoid contaminating sterile media, tips and cultures while you work.

## Where you work

- **Biosafety cabinet (BSC).** Cell culture and work with many microorganisms is done in a Class II BSC, which draws air through HEPA filters to protect the work, you and the room. Do not block the front or rear grilles, work at least several centimetres inside the cabinet, and move slowly so you do not disrupt the air curtain. Bunsen burners are generally **not** used in a BSC: the heat disrupts airflow and can damage the filter.
- **Open bench with a flame.** Many microbiology teaching labs work on the bench beside a Bunsen burner, which creates an updraft, and flame loops and tube mouths. Follow what your lab's SOP specifies.

The biosafety level of the agent (BSL-1 to BSL-4) decides which containment and practices are required. See the CDC/NIH guide *Biosafety in Microbiological and Biomedical Laboratories (BMBL)*: https://www.cdc.gov/labs/bmbl/ (Checked on 2026-10-23.)

## The core habits

1. **Prepare.** Plan the steps, gather everything you need, and label tubes and plates before you start, so the cabinet is not crowded and open containers wait as little as possible.
2. **Hands and PPE.** Wash hands, put on a clean lab coat and gloves as your SOP requires. Spray gloves with 70 percent ethanol and let them dry.
3. **Disinfect the work surface** before and after, usually with 70 percent ethanol (which works best with some water present and needs contact time) or another approved disinfectant.
4. **Disinfect items going into the cabinet** by wiping or spraying bottles and racks.
5. **Open containers only when needed and close them promptly.** Hold caps in your hand or place them face down on a disinfected surface, as your SOP specifies. Never touch the inside of a cap, the neck of a bottle or a sterile tip.
6. **Never pass hands or arms over open containers.** Particles fall from you onto whatever is below.
7. **Use each sterile pipette or tip once.** If it touches anything non-sterile, discard it and take a new one.
8. **Work in a logical flow**, from clean to dirty, and keep waste at the side.
9. **Clean up.** Discard waste into the correct biohazard containers, disinfect the surface again, remove gloves and wash hands.

## Worked example: an aseptic transfer of medium

In a BSC, a technician moves 10 mL of sterile medium into a labelled sterile flask:

1. Wipes the medium bottle and flask with 70 percent ethanol and places them in the cabinet.
2. Attaches a sterile 10 mL serological pipette to the pipette controller without touching the tip.
3. Opens the bottle, keeps the cap in the other hand, draws 10 mL, recaps the bottle.
4. Opens the flask, dispenses down the inside wall without touching the neck, recaps the flask.
5. Discards the pipette into the waste container without touching anything else on the way.

Every step keeps sterile surfaces from touching non-sterile ones and keeps containers open as briefly as possible.

## Checking your technique

- **Negative controls:** incubate a tube or plate of medium you opened but did not inoculate. Growth means your technique or materials introduced contamination.
- **Settle plates and environmental monitoring** are used in cleanrooms and biomanufacturing.
- In cell culture, watch for cloudy medium, sudden colour changes and floating particles.

## Common mistakes

- Talking, coughing or reaching over open containers.
- Overcrowding the cabinet and blocking its grilles.
- Touching the tip of a pipette to the outside of a bottle neck and carrying on.
- Leaving bottles open while you fetch something.
- Skipping the negative control.

## Quick recap

- Sterile is a state; aseptic is a set of working habits.
- Work in a BSC (no flame) or beside a flame, as your SOP says.
- Disinfect, open briefly, never touch sterile surfaces, never reach over open containers, single-use tips.
- Prove your technique with negative controls.

Aseptic work with real cultures is learned only in your teaching lab or workplace, after biosafety training and under supervision.

## Try it

Open the **Do** tab to rehearse the motions of an aseptic transfer using only clean, empty containers and tap water. No microbes, media or cultures are involved.


Understand the habits of aseptic transfer that keep cultures and reagents free of contamination.

Aseptic technique

Aseptic Technique

Sterile Technique

Calibrate and log instruments, compare fluorescence and electron microscopy, and practise the habits of aseptic technique.

Lab Techniques & Instrumentation

About this course

Course content

Core Bench Skills

Separation and Analysis

Instruments and Upkeep

Where this course leads

Next course on the path

Careers that use it