Extract nucleic acids, run PCR and qPCR, read gels, Western blots and ELISAs, compare sequencing methods, and work with sequences and BLAST.

Molecular Biology Methods

Download a spin-column kit handbook and find the protocol for one sample type (for example, cultured cells or blood).

Read the whole protocol once without writing anything.

On a printout or in a notebook, label each step as lyse, bind, wash, dry or elute.

Mark every step that needs preparation before the day (buffer reconstitution, pre-heating, thawing).

Circle each point where you keep the flow-through versus discard it, and each safe stopping point.

Write down where an extraction blank fits in and how you would label your tubes (sample ID, date, initials).

Calculate concentration, A260/A280 and A260/A230 for three DNA readings: (a) A260 0.84, A280 0.46, A230 0.41; (b) A260 0.50, A280 0.33, A230 0.48; (c) A260 0.30, A280 0.16, A230 0.12. Decide which extracts you would use.

If your teaching lab runs an extraction practical, compare your annotated protocol with the one your instructor uses and ask about any differences.

## Why it matters

Almost every molecular method in this course starts with clean nucleic acid. PCR, qPCR, sequencing and cloning all fail, or quietly give wrong answers, when the DNA or RNA going in is degraded, contaminated or too dilute. A biological technician may run dozens of extractions a day, and a medical laboratory technologist may run them on automated platforms for infectious disease or genetic tests. In every case the job is the same: get the nucleic acid out of the cells, away from everything else, and prove it is good enough to use.

## The core idea: lyse, bind, wash, elute

Most modern extractions use a **silica spin column** or **magnetic silica beads**. The steps are the same in almost every kit:

1. **Lyse.** Break open cells and nuclei with a lysis buffer (detergent plus salts), often with an enzyme such as proteinase K to digest proteins. Tough samples (plant tissue, bacteria with thick walls) may need mechanical disruption too.
2. **Bind.** Add a high concentration of a chaotropic salt (and usually ethanol). Under these conditions nucleic acids stick to silica, while most proteins and other debris do not.
3. **Wash.** Spin wash buffers (usually ethanol-based) through the column to remove salts and leftover contaminants. A final "dry spin" removes residual ethanol, which would inhibit later reactions.
4. **Elute.** Add a small volume of low-salt buffer or nuclease-free water. Nucleic acid releases from the silica and collects in a clean tube.

Older methods, such as phenol-chloroform extraction (and its RNA cousin, TRIzol), separate nucleic acids by partitioning into a water layer. They work well but use hazardous solvents, so they are done only in a fume hood after specific training.

## DNA versus RNA

RNA is far less forgiving than DNA:

- **RNases are everywhere** (skin, dust, many reagents) and do not need cofactors. RNA work uses RNase-free tubes and tips, fresh gloves, a clean bench and often a dedicated area.
- **Work fast and cold.** Samples are kept on ice or put into a stabilising reagent immediately.
- **Remove DNA.** RNA preps for gene expression are usually treated with **DNase** so genomic DNA does not show up in later qPCR.

## Checking what you got

You always measure yield and purity before using an extract, usually on a microvolume UV spectrophotometer:

| Measurement | What it tells you | Typical target |
|---|---|---|
| A260 | Concentration. An A260 of 1.0 is about 50 ng/uL for double-stranded DNA and about 40 ng/uL for RNA | Depends on the method downstream |
| A260/A280 | Protein contamination | about 1.8 for DNA, about 2.0 for RNA |
| A260/A230 | Salt, chaotrope or phenol carryover | about 2.0 to 2.2 |

**Worked example.** A DNA eluate (undiluted, 1 cm path equivalent) reads A260 = 0.84, A280 = 0.46, A230 = 0.41.

- Concentration = 0.84 x 50 ng/uL = **42 ng/uL**.
- A260/A280 = 0.84 / 0.46 = **1.83**: little protein.
- A260/A230 = 0.84 / 0.41 = **2.05**: little salt carryover.

This extract looks usable. If A260/A230 had been 0.9, you would suspect chaotropic salt carryover, often from a skipped dry spin or wash, and expect PCR to struggle.

For RNA, purity ratios are not enough: integrity matters too. Labs check it on an automated electrophoresis instrument that gives an **RNA integrity number (RIN)** from 1 (degraded) to 10 (intact).

## Following a protocol

A kit protocol is a controlled procedure, not a suggestion. Before you start, read it once end to end, and mark:

- **Steps you must prepare in advance**, such as adding ethanol to a wash buffer concentrate or pre-heating a block.
- **Steps that are easy to get wrong**, such as which wash buffer goes first or whether to discard the flow-through.
- **Stopping points**, where the sample can be safely stored.
- **Your controls**: an extraction blank (no sample) shows whether reagents or technique introduced contamination.

Record lot numbers, the protocol version and any deviation in your notebook, as you learned in the lab data course.

## Safety and real-world training

Real samples can be infectious (clinical specimens, cultured pathogens) and lysis reagents are often irritant or toxic. Extraction is done only in a teaching or employer lab, at the right biosafety level, after institution training and under supervision. The CDC's *Biosafety in Microbiological and Biomedical Laboratories* (BMBL) explains biosafety levels: https://www.cdc.gov/labs/bmbl/index.html (Checked on 2026-10-23.) Nothing in this course asks you to extract nucleic acids at home.

## Common mistakes

- Skipping the dry spin, so ethanol carries over into the eluate.
- Mixing up flow-through and column: discarding the column instead of the flow-through.
- Overloading the column with too much starting material, which lowers purity and yield.
- Touching tube rims and tips during RNA work.
- Trusting A260 alone without checking the ratios.

## Quick recap

- Silica methods: lyse, bind (chaotropic salt), wash (ethanol), dry, elute (low salt).
- RNA needs RNase-free technique, speed, cold and usually DNase treatment.
- Check concentration (A260) and purity (A260/A280 about 1.8 DNA or 2.0 RNA; A260/A230 about 2.0 to 2.2).
- Run an extraction blank and record lots and deviations.

## Try it

Open the **Do** tab to annotate a real kit protocol and judge three sets of spectrophotometer readings, on paper.


Follow the logic of a DNA or RNA extraction protocol and judge the yield and purity of what you get.

DNA/RNA extraction

DNA extraction explained | biology animation

How to isolate RNA from tissue or cells

## Why it matters

The polymerase chain reaction (PCR) makes millions of copies of one short stretch of DNA in about an hour. It is behind genotyping, cloning, pathogen detection, forensic profiling and most sequencing library preparation. Biological technicians set up PCRs constantly, and most failures trace back to two things: badly designed primers and careless reaction setup.

## The core idea

PCR repeats three temperature steps, usually 25 to 35 times, in a thermal cycler:

1. **Denature** at about 94 to 98 C: the two DNA strands separate.
2. **Anneal** at about 50 to 65 C: two short **primers** bind to their complementary sites on opposite strands, flanking the target.
3. **Extend** at about 72 C: a heat-stable DNA polymerase (such as Taq) adds nucleotides to the 3' end of each primer.

Each cycle roughly doubles the target, so after n perfect cycles one copy becomes 2^n copies. In practice efficiency is below 100 percent and the reaction plateaus as reagents run out, which is why plain PCR is good for "is it there, and how big is it?" but not for measuring how much was there to begin with. That is what qPCR (next lesson) is for.

## What goes in the tube

A typical 25 uL reaction contains:

| Component | Role |
|---|---|
| Template DNA | The DNA containing your target (often 1 to 100 ng genomic DNA) |
| Forward and reverse primers | Define the start and end of the product (often 0.1 to 0.5 uM each) |
| dNTPs | The A, C, G, T building blocks |
| Polymerase buffer with Mg2+ | Magnesium is a required cofactor; its concentration affects specificity |
| DNA polymerase | Copies the DNA |
| Nuclease-free water | Brings the volume up |

Labs usually make a **master mix** of everything except template, split it into tubes, then add template last. Always include a **no-template control (NTC)**, with water instead of template, to catch contamination, and a **positive control** known to work.

## Designing primers

Primers are short single-stranded DNA pieces. Common rules of thumb:

- **Length** 18 to 25 nucleotides.
- **GC content** 40 to 60 percent.
- **Melting temperature (Tm)** roughly 52 to 65 C, with the two primers **within about 5 C** of each other.
- A G or C in the last one or two bases at the 3' end (a "GC clamp") helps the polymerase start, but avoid more than three G or C in the last five bases.
- Avoid runs of four or more of one base, and avoid primers that pair with themselves or each other (hairpins and primer dimers).
- The reverse primer is written 5' to 3' and matches the **reverse complement** of the top strand.

A quick Tm estimate for short primers is the **Wallace rule**:

```text
Tm = 2 x (number of A + T) + 4 x (number of G + C)
```

**Worked example.** Primer `AGCTGACCTGAAGGCTCTTG` has 20 bases: A 4, T 5, G 6, C 5. A + T = 9 and G + C = 11, so Tm = 2 x 9 + 4 x 11 = **62 C**. GC content is 11 / 20 = **55 percent**. Both are in range.

The Wallace rule is only a rough guide (it ignores salt and the order of bases). Real design uses nearest-neighbour calculators such as Primer-BLAST from NCBI (https://www.ncbi.nlm.nih.gov/tools/primer-blast/, checked on 2026-10-23), which also checks that the primers match only your intended target. A common starting annealing temperature is about 3 to 5 C below the lower primer Tm, refined with a gradient PCR.

## Reading the result

PCR products are usually checked on an agarose gel (module 2). You expect one sharp band at the predicted size, nothing in the NTC, and a band in the positive control. Extra bands suggest non-specific priming (try a higher annealing temperature); a smear suggests too much template or too many cycles; a band in the NTC means contamination, so the whole run is suspect.

## Avoiding contamination

Because PCR is so sensitive, one stray product molecule can give a false positive. Good labs separate **pre-PCR** (reagent and setup) and **post-PCR** (gel) areas, use filter tips, never bring opened products back to the setup area, and clean benches before setup. Clinical molecular labs follow these rules strictly and are inspected on them.

## Common mistakes

- Forgetting the NTC, so contamination goes unnoticed.
- Writing the reverse primer as the top-strand sequence instead of its reverse complement.
- Primer Tm values 10 C apart, so one primer barely binds at the chosen temperature.
- Adding template to the master mix before splitting it.
- Opening post-PCR tubes in the setup area.

## Quick recap

- Denature, anneal, extend; each cycle roughly doubles the target.
- Master mix, template last, NTC and positive control every time.
- Primers: 18 to 25 nt, 40 to 60 percent GC, Tm within 5 C, no runs or dimers.
- Wallace rule Tm = 2(A+T) + 4(G+C) is a quick estimate; use Primer-BLAST for real design.

## Try it

Open the **Do** tab. A Python primer checker is ready except for the Tm formula, which you write.


Understand how PCR copies a target and check a primer pair before you set up a reaction.

PCR (Polymerase Chain Reaction)

How to Design Primers for PCR

## Why it matters

Quantitative PCR (qPCR, also called real-time PCR) measures how much of a target sequence was in a sample, not just whether it was there. Research labs use it to compare gene expression between treated and untreated cells. Clinical labs use it to detect and quantify pathogens (viral load, for example). If you work in a molecular lab, you will read Ct values and fold changes every week.

## The core idea: watch the reaction as it happens

A qPCR instrument is a thermal cycler that measures fluorescence in each well after every cycle. The fluorescence comes from either:

- a **DNA-binding dye** such as SYBR Green, which glows when bound to double-stranded DNA, or
- a **probe** (for example a hydrolysis or "TaqMan" probe), which glows only when the specific target is copied.

Plotting fluorescence against cycle number gives an S-shaped **amplification curve**. The software sets a **threshold** in the exponential part of the curve. The **Ct** (cycle threshold, also called Cq) is the cycle at which a well's curve crosses it.

The key fact: **more starting template means a lower Ct**. If efficiency is near 100 percent, the target doubles each cycle, so a sample with twice as much template crosses one cycle earlier. A difference of 3.3 cycles is about a 10-fold difference.

## Relative expression: the delta-delta-Ct method

To compare expression of a **target gene** between a **treated** and a **control** sample, you also measure a **reference gene** (for example GAPDH or ACTB) whose expression should not change. It corrects for differences in how much RNA went into each reaction. The Livak method:

1. **Delta Ct** for each sample: Ct(target) - Ct(reference).
2. **Delta-delta Ct**: delta Ct(treated) - delta Ct(control).
3. **Fold change** = 2^(-delta-delta Ct).

**Worked example.** Means of triplicate wells:

| Sample | Ct target | Ct reference | Delta Ct |
|---|---|---|---|
| Control | 25.0 | 18.0 | 7.0 |
| Treated | 23.0 | 18.2 | 4.8 |

Delta-delta Ct = 4.8 - 7.0 = **-2.2**. Fold change = 2^2.2 = **4.6**. The target is about 4.6 times higher in treated cells. A positive delta-delta Ct gives a fold change below 1, meaning lower expression.

## When the method is valid

The 2^-delta-delta-Ct formula assumes:

- **Both assays have efficiency close to 100 percent** (about 90 to 110 percent), checked with a dilution series standard curve. If they differ, use an efficiency-corrected method (such as Pfaffl).
- **The reference gene really is stable** under your conditions. Many labs validate two or three reference genes.
- **Ct values are in a reliable range.** Very late Ct values (often above about 35) are noisy and may be non-specific.

Good practice is summarised in the **MIQE guidelines** (Bustin and colleagues, *Clinical Chemistry*, 2009, https://doi.org/10.1373/clinchem.2008.112797, checked on 2026-10-23), which list what a qPCR experiment must report.

## Controls and quality checks

- **No-template control (NTC):** should show no amplification, or a Ct far later than any sample.
- **No-reverse-transcriptase control (no-RT):** for RNA work, shows whether genomic DNA is giving signal.
- **Technical replicates:** usually triplicates. Replicate Ct values more than about 0.3 to 0.5 cycles apart suggest pipetting problems.
- **Melt curve (dye-based assays):** one clean peak means one product; extra peaks mean primer dimers or off-target products.

## Absolute quantification

Clinical and environmental labs often need actual copy numbers. They run a **standard curve** of known copy numbers and read unknowns off it. The slope of Ct against log10(copies) gives efficiency: E = 10^(-1/slope) - 1. A slope of -3.32 means 100 percent efficiency.

## Common mistakes

- Subtracting in the wrong order, so the fold change is inverted.
- Averaging fold changes instead of averaging Ct values first.
- Using a reference gene that the treatment actually changes.
- Ignoring a positive NTC or no-RT control.
- Reporting fold changes from Ct values near 38.

## Quick recap

- Lower Ct = more starting target; about 1 cycle = 2-fold at 100 percent efficiency.
- Delta Ct = target - reference; delta-delta Ct = treated - control; fold = 2^(-delta-delta Ct).
- Check efficiency, reference stability, replicates, NTC, no-RT and melt curves.
- Follow MIQE when you report results.

## Try it

Open the **Do** tab. A Python function computes delta Ct for you; you write the delta-delta-Ct and fold-change line.


Read Ct values from quantitative PCR and compute relative gene expression with the delta-delta-Ct method.

qPCR

qPCR and PCR Explained: Real-Time PCR Animation

How To Perform The Delta-Delta Ct Method (In Excel)

Follow a DNA or RNA extraction protocol, set up a PCR with a checked primer pair, and compute relative expression from qPCR with delta-delta-Ct.