Biochemical analysis is a cornerstone of applied biology, allowing professionals to identify, separate, and characterize biomolecules such as proteins, enzymes, nucleic acids, and lipids. In Kenyan laboratories, techniques like tissue homogenization and ultracentrifugation are routinely used to isolate cellular components for research, diagnostics, and industrial applications. Understanding enzymes, their classifications and properties, is fundamental, as these biological catalysts drive the reactions necessary for biomolecule separation and qualitative analysis. Mastery of enzyme concepts equips applied biology graduates for roles in hospitals, agricultural research centers, and biotechnology firms, where accurate biochemical assays can impact everything from disease diagnosis to food quality testing.

2.1 Biomolecule separation procedures

Understanding the mechanisms of action in biochemical analysis is vital for Applied Biology professionals in Kenya, especially when handling tissue samples or analyzing biomolecules in clinical, agricultural, and research settings. The mechanisms by which cellular components are separated and identified underpin much of modern laboratory diagnostics, from disease screening in county hospitals to quality assurance in food processing plants. These processes rely on physical and chemical principles to disrupt cells, isolate organelles, and observe molecular behaviour, ensuring that accurate and reliable results are achieved.

Cell Lysis and Buffering

Cell lysis is the foundational step in biomolecule separation, allowing access to intracellular components by breaking cell membranes. In Kenya, this technique is routinely employed in medical laboratories, agricultural research stations, and university teaching labs, where tissue samples from plants, animals, or microorganisms are processed for further analysis. The use of buffer solutions during lysis is crucial to maintain the integrity of the sample and prevent unwanted chemical reactions.

Principles of Cell Lysis

• Mechanical Disruption: Physical force, such as grinding with a blender or mortar and pestle, breaks open cell membranes. This is especially effective for tough plant tissues or animal organs, as seen in county hospital pathology labs handling biopsy samples.
• Buffer Solution Use: A dense, inert aqueous solution, commonly sucrose or brine, suspends the sample. This prevents osmotic shock and unwanted reactions, preserving organelle structure for later analysis.
• Porcelain Pores and Container Wall: Grinding against the container's wall and through porcelain pores creates tiny fragments, facilitating thorough lysis and release of organelles.
• Partial Lysis: Not all cells or organelles are fully lysed; some remain intact or are damaged, which must be addressed during subsequent centrifugation steps.
• Sample Suspension: The resulting liquid contains a mix of intact cells, lysed cells, and free organelles, ready for further separation.

Lysis Buffer and Other Lysis Methods

A lysis buffer does more than suspend the sample. It holds the pH steady (commonly near pH 7.4 for animal tissue) so that proteins and enzymes keep their shape, and it supplies salts at a concentration close to that inside the cell so that organelles do not swell or shrink. Many lysis buffers also contain extra agents: a mild detergent to dissolve membranes, EDTA to bind metal ions that some degrading enzymes need, and protease inhibitors that protect proteins from breaking down. Example: a university teaching laboratory preparing a liver homogenate would grind the tissue in ice-cold buffered sucrose so that the mitochondria stay intact.

• Homogenising: A glass or Potter-type homogeniser shears soft animal tissue gently, which suits the isolation of intact organelles.
• Grinding with sand: A mortar and pestle with clean sand breaks tough plant cell walls, as when a seed or leaf extract is prepared in a school laboratory.
• Sonication: High-frequency sound waves burst bacterial and other small cells; the tube is kept in ice because the process heats the sample.
• Detergent lysis: Detergents such as SDS or Triton X-100 dissolve the membrane lipids and are used when the organelles do not need to stay whole.
• Freeze-thaw cycles: Repeated freezing and thawing forms ice crystals that rupture the membranes of cultured cells and microorganisms.

Typical Mistakes in Cell Lysis

• Insufficient Grinding: Leads to incomplete lysis, reducing yield of desired biomolecules.
• Incorrect Buffer Concentration: Can cause osmotic damage or loss of sample integrity, affecting downstream results.
• Overheating During Grinding: Excessive heat may denature proteins and enzymes, compromising analysis accuracy.
• Contamination: Improper cleaning of equipment introduces unwanted substances, skewing results.
• Delayed Processing: Allowing samples to sit after lysis increases risk of degradation by endogenous enzymes.

Ultracentrifugation: Physical Separation of Biomolecules

Ultracentrifugation is a powerful technique used to separate cellular components based on their size, density, and shape. This method is widely applied in Kenyan research institutes and clinical labs for isolating organelles, viruses, and macromolecules from complex biological samples. The process relies on high-speed rotation to create intense centrifugal forces, driving sedimentation of particles. Spinning the lysate in a series of steps at increasing speed, so that each step removes one class of particle, is called differential centrifugation.

Steps in Ultracentrifugation

1. Preparation of Lysate: The tissue sample is first lysed and suspended in buffer, ensuring a uniform mixture of cellular fragments.
2. Loading the Ultracentrifuge: Samples are placed in tubes and loaded into the rotor of the ultracentrifuge, which is maintained at low temperature and pressure.
3. Initial Centrifugation: The lysate is spun at low speed, causing larger fragments such as nuclei to sediment and form a pellet.
4. Removal of Pellet: The pellet is carefully removed, and the supernatant is retained for further centrifugation.
5. Sequential Centrifugation: The supernatant is subjected to progressively higher speeds, sedimenting smaller organelles like mitochondria, lysosomes, and ribosomes.
6. Layer Extraction: Each pellet is extracted after each centrifugation step, resulting in distinct layers containing different cellular components.

Factors Affecting Sedimentation

• Mass and Shape: Larger, more massive molecules sediment faster under centrifugal force.
• Partial Specific Volume: The volume occupied by a molecule influences its sedimentation velocity.
• Solvent Density: Higher density solvents slow sedimentation, while lower density solvents accelerate it.
• Rotor Size and Speed: Larger rotors and higher speeds generate greater centrifugal force, enhancing separation.
• Temperature: Maintaining low temperatures prevents sample degradation and ensures accurate results.

Precautions in Centrifugation

• Balance the tubes: Opposite tubes must have the same mass, checked on a balance, because an unbalanced rotor vibrates and can fail violently at high speed.
• Use the correct rotor and tubes: Each rotor and tube type is rated for a maximum speed; exceeding it can crack the tubes or damage the rotor.
• Keep the sample cold: Ice-cold buffer and a refrigerated rotor slow the enzymes that would otherwise break down the proteins.
• Label every fraction: Pellets and supernatants are labelled at once and stored cold so that the fractions are not confused.
• Wait for the rotor to stop: The lid is opened only after the rotor has stopped completely, and spills are cleaned with the disinfectant the laboratory prescribes.

Analytical Ultracentrifuges and Optical Systems

Analytical ultracentrifuges are specialized instruments used to study the behaviour of biomolecules during centrifugation. Their application is common in university research labs and biotechnology companies in Kenya, where scientists analyze protein complexes, nucleic acids, and other macromolecules.

Name	Specification	Use
Analytical Ultracentrifuge	Up to 500,000 g; optical detection systems (light absorption, Schlieren, Rayleigh interferometric)	Studying sedimentation behaviour and molecular weight determination

Types of Optical Detection Systems

• Light Absorption System: Measures absorbance of light by sample, useful for quantifying protein concentration in pharmaceutical labs.
• Schlieren System: Detects changes in refractive index, allowing visualization of concentration gradients in biological samples.
• Rayleigh Interferometric System: Measures interference patterns to determine molecular mass and interactions.
• Real-Time Monitoring: Enables observation of sedimentation velocity, facilitating calculation of sedimentation coefficients.
• Data Analysis: Optical data is used to estimate molecular weight, size, and shape, informing further biochemical analysis.

Sedimentation Coefficient and Molecular Weight Determination

The sedimentation coefficient (S) is a critical parameter in ultracentrifugation, reflecting how rapidly a molecule sediments under centrifugal force. Calculating S provides insight into the molecular weight and structural properties of biomolecules, which is essential for Applied Biology professionals working in diagnostic and research settings. It is expressed in Svedberg units (1 Svedberg = 10⁻¹³ seconds), so a larger S value means faster sedimentation.

Components Influencing Sedimentation Coefficient

• Particle Mass: Heavier particles have larger S values, indicating faster sedimentation rates.
• Particle Shape: Spherical particles sediment more efficiently than elongated or irregular shapes.
• Solvent Viscosity: Higher viscosity slows sedimentation, reducing S value.
• Temperature: Elevated temperatures can alter viscosity and molecular behaviour, affecting S.
• Rotor Speed: Increased speed enhances centrifugal force, increasing sedimentation velocity and S value.

2.1.1 Iodine/iodide test (starch test)

The iodine test shows whether a sample contains starch, the storage carbohydrate of plants. Starch is a mixture of amylose and amylopectin, and the test is used on foods such as maize flour, cassava, rice and potato, and on leaves to show that a plant has been photosynthesising. Example: a quality officer at a maize milling company can use it to find out whether a bag of flour labelled as pure maize meal has been bulked with another starch-containing filler, although the test only shows that starch is present, not which plant it came from.

Principle

Iodine solution contains iodine (I₂) dissolved in potassium iodide (KI), which together form the triiodide ion (I₃⁻). Amylose is a long coiled chain of glucose units, and the triiodide ions slide into the centre of the coil. The trapped iodine chain absorbs light strongly and gives an intense blue-black colour. Amylopectin is branched and has only short coiled stretches, so it gives a weaker red-purple or red-brown colour, and glycogen gives a red-brown colour. Glucose, sucrose, cellulose and protein do not form the complex, so the iodine stays yellow-brown.

Reagents and Equipment

Name	Specification	Use
Iodine solution	Iodine in potassium iodide, about 0.01 mol/dm³ iodine (Lugol's iodine is stronger, about 1% iodine in 2% KI)	Reacts with starch to give blue-black colour
Test tubes and rack	Clean, labelled, 10 to 15 cm³	Hold the sample and controls
Dropper or dropping pipette	Plastic, 1 cm³	Adds the iodine drop by drop
Spotting tile or white tile	White ceramic	Shows the colour on solid foods
Starch solution, 1%	1 g soluble starch in 100 cm³ hot water, cooled	Positive control
Distilled water	Clean	Negative control

Procedure

1. Put on the laboratory coat, gloves and goggles, and label three clean test tubes: sample, positive control and negative control.
2. Place about 2 cm³ of the liquid sample in the first tube. For a solid food, grind a small amount with a little water, or place a small piece on a white tile.
3. Place about 2 cm³ of 1% starch solution in the second tube and 2 cm³ of distilled water in the third.
4. Make sure all the liquids are cool. Hot solutions must be cooled to room temperature first because heat destroys the colour.
5. Add 2 to 3 drops of iodine solution to each tube, or to the solid food on the tile. Shake the tubes gently to mix.
6. Observe the colour at once and again after about one minute against a white background.
7. Record the colour for each tube in a results table and compare the sample with the two controls.

Results and Interpretation

Observation	Meaning
Yellow-brown stays unchanged	Starch is absent
Blue-black	Starch (amylose) is present
Red-purple or red-brown	Amylopectin, glycogen or dextrins (partly broken down starch) are present
Blue-black colour fades on warming and returns on cooling	Confirms starch, because the complex breaks on heating and re-forms when cool

The starch control must turn blue-black and the water control must stay yellow-brown. If either control fails, the reagent is faulty and the test is repeated with fresh solution.

Safety

• Iodine stains skin, clothing and work surfaces brown, so wear gloves and a laboratory coat.
• Iodine solution is harmful if swallowed and its vapour irritates the eyes, so keep bottles closed and wear goggles.
• Never taste or eat any food that has been used in a laboratory test.
• Wash hands after the practical and dispose of the solutions as the laboratory instructs.
• Keep the iodine bottle away from direct sunlight, which breaks down the solution.

Sources of Error

• Testing a hot sample: The blue-black complex does not form at high temperatures, giving a false negative.
• Too much iodine: A large excess gives a dark brown colour that hides a weak positive.
• Strongly acidic or alkaline samples: Extreme pH changes the iodine and can destroy the colour, so very acidic samples such as lemon juice should be neutralised first.
• Old iodine solution: Iodine is lost by evaporation and light, so stale reagent gives weak or no colour.
• Coloured samples: Dark foods hide the colour change; diluting the sample or using a white tile helps.

2.1.2 Reducing sugar test (Benedict's test)

Benedict's test detects reducing sugars, which are sugars that can donate electrons to another substance. All monosaccharides (glucose, fructose, galactose) and some disaccharides (maltose, lactose) are reducing sugars. Example: a county referral hospital laboratory once used Benedict's solution to screen urine for glucose, and a school laboratory uses it to compare the sugar in ripe and unripe bananas.

Principle

In hot alkaline solution, the free aldehyde group of an aldose (or the ketose, which converts to an aldose form in alkali) reduces blue copper(II) ions (Cu²⁺) in the reagent to copper(I) ions, which form insoluble copper(I) oxide (Cu₂O). The precipitate is brick red. The sodium citrate in Benedict's solution keeps the copper(II) ions dissolved in the alkaline sodium carbonate solution so that they do not precipitate as copper hydroxide. The more reducing sugar present, the more copper(I) oxide forms and the further the colour moves from blue to red. Sucrose has no free aldehyde or ketone group because its two sugar units are joined through both reducing groups, so it gives no reaction until it is hydrolysed.

Reagents and Equipment

Name	Specification	Use
Benedict's solution	Per litre: 17.3 g copper(II) sulphate pentahydrate, 173 g sodium citrate, 100 g anhydrous sodium carbonate; clear blue	Gives a coloured response to reducing sugar
Test tubes and rack	Heat-resistant, labelled	Hold the reaction mixture
Water bath	Beaker of boiling water on a hot plate or burner, about 95 to 100 °C	Heats the tubes evenly without direct flame
Test tube holder and marker	Wooden or metal	Safe handling and labelling
Glucose solution, 1%	1 g glucose in 100 cm³ water	Positive control
Distilled water	Clean	Negative control
Dilute hydrochloric acid and sodium hydrogencarbonate	About 1 mol/dm³ acid; solid or solution base	Hydrolyse and then neutralise sucrose for the non-reducing sugar test

Procedure

1. Wear the laboratory coat, gloves and goggles, and label three tubes: sample, positive control (glucose) and negative control (distilled water).
2. Place 2 cm³ of the sample in the first tube, 2 cm³ of glucose solution in the second and 2 cm³ of distilled water in the third. A solid food is first ground in water and filtered.
3. Add 2 cm³ of Benedict's solution to each tube and mix by gentle shaking.
4. Put all three tubes in the boiling water bath at the same time and heat for 3 to 5 minutes. Point the mouths of the tubes away from people.
5. Remove the tubes with a holder and stand them in the rack. Do not shake while hot.
6. Observe the colour and any precipitate, and compare it with the controls and the colour chart.
7. Record the result as negative, trace, low, moderate, high or very high (green to brick red).
8. For a negative sample, test for a non-reducing sugar: boil a fresh 2 cm³ of sample with 2 cm³ of dilute hydrochloric acid for about 2 minutes, cool, add small amounts of sodium hydrogencarbonate until fizzing stops (the solution is then neutral or slightly alkaline), and repeat steps 3 to 7. A brick-red colour now shows that sucrose was present and has been hydrolysed to glucose and fructose.

Results and Interpretation

Colour after heating	Amount of reducing sugar
Blue, no change	None
Green, clear or cloudy	Trace to low (about 0.5 g per 100 cm³)
Yellow to orange	Moderate
Orange-red	High
Brick-red precipitate	Very high (above about 2 g per 100 cm³)

The sequence from blue to green, yellow, orange and brick red shows an increasing amount of reducing sugar. The test is semi-quantitative: it gives a rough amount, not an exact figure. Uncooked glucose or fructose controls give the expected colours.

Safety

• Hot water and boiling liquids can scald, so use a holder, never hold a tube with the fingers, and point tubes away from faces.
• Benedict's solution is irritant and harmful to the eyes, and copper salts are toxic to aquatic life, so rinse spills with plenty of water and dispose of waste as instructed.
• Dilute hydrochloric acid is corrosive: wear goggles and wash splashes from the skin at once.
• Sodium hydrogencarbonate fizzes when added to acid, so add it slowly to avoid overflow.
• Heat the tubes in a water bath rather than a direct flame, because direct heating makes the liquid spit.

Sources of Error

• Too short heating or a water bath that is not boiling: The reaction is slow at low temperature and the colour is under-read.
• Too much sample: The colour of the sample hides the copper colour, or the blue reagent is used up and the test seems weaker than it is.
• Other reducing substances: Ascorbic acid, some drugs and some preservatives also reduce copper(II) and give a false positive.
• Incomplete neutralisation after hydrolysis: Acid left in the solution prevents the reaction, giving a false negative for sucrose.
• Reading too late: The precipitate settles on standing, so the colour should be read at once, with the tube still.

2.1.3 Translucent test (lipids)

The translucent test, also called the grease spot test, is the simplest way to show that a sample contains lipids (fats and oils). Lipids are not soluble in water and do not evaporate, so they leave a permanent mark on paper that water does not. Example: a farm laboratory checking nuts or seeds for oil content, or a hotel kitchen checking whether a pastry has been made with butter, can use it as a quick screening step before a laboratory analysis.

Principle

Paper is made of cellulose fibres, which scatter light and so look opaque. When a lipid soaks into the paper, it fills the air spaces between the fibres. Lipid has a refractive index closer to that of cellulose than air has, so light passes through the paper instead of being scattered, and the spot appears translucent (see-through). Water soaks in as well, but it evaporates on drying, and the paper returns to its opaque state. A spot that is still translucent after the paper is dry therefore shows that a non-volatile, water-insoluble substance, a lipid, is present.

Reagents and Equipment

Name	Specification	Use
Filter paper or brown (unglazed) paper	White filter paper or plain brown paper, 5 cm square	Receives the sample and shows the spot
Glass rod or spatula	Clean	Spreads the sample
Mortar and pestle	Porcelain	Crushes solid foods such as nuts or seeds
Cooking oil	Vegetable oil	Positive control
Distilled water	Clean	Negative control
Light source	Window or lamp	Shows the translucent spot

Procedure

1. Label three pieces of paper: sample, oil (positive control) and water (negative control).
2. Crush a solid sample into a paste, or take a drop of a liquid sample.
3. Rub or place a small amount of the sample on the first piece of paper. Place one drop of oil on the second and one drop of water on the third.
4. Leave the papers to dry. A warm place or a short time in a drying oven at a low setting speeds up drying, but avoid overheating.
5. When the papers are completely dry, hold each one up to the light.
6. Observe whether a see-through spot is visible and compare it with the two controls.
7. Record the result and, if needed, repeat with a second sample of the same food to confirm.

Results and Interpretation

• Positive result: A translucent, greasy spot remains after drying, which shows that lipids are present.
• Negative result: No spot remains after the paper has dried, meaning lipids are absent or present in very small amounts.
• Oil control: The spot stays translucent and does not dry out, confirming that the paper and method work.
• Water control: The spot disappears on drying, confirming that the paper is not itself translucent.
• Strength of the spot: A larger or clearer spot suggests more lipid, but the test is only qualitative and cannot give a percentage.

Safety

• Do not eat or taste any food tested in the laboratory.
• Keep the drying oven low and away from flammable liquids if oil-soaked paper is dried with heat.
• Wash hands with soap afterwards, because oils spread onto benches and handles.
• Dispose of oil-soaked paper in the waste bin and not in a sink.
• If an ethanol emulsion test is used as a second confirmation, work away from flames because ethanol is flammable.

Sources of Error

• Testing before the paper is dry: A wet spot from water looks translucent and gives a false positive.
• Using glossy or coated paper: The coating does not absorb the sample, so no spot forms and the result may be missed.
• Very small lipid amounts: Lean foods give a faint spot that is hard to see, producing a false negative.
• Volatile oils: Essential oils such as those from some plants evaporate and leave no spot.
• Greasy hands or dirty equipment: Handling the paper with oily fingers produces a spot that is not from the sample.

2.1.4 Biuret test (proteins)

The Biuret test shows whether a sample contains protein by detecting peptide bonds. It is used to screen foods such as milk, eggs, beans and meat, and in clinical laboratories for total protein in blood serum. Example: a dairy cooperative laboratory can use it to confirm that milk samples contain protein before more detailed tests, and a school laboratory uses it to compare egg white with beans.

Principle

In alkaline solution, copper(II) ions form a coordination complex with the nitrogen atoms of peptide bonds (the CO-NH links between amino acids). The complex is violet or purple. A molecule needs at least two peptide bonds, that is, at least three amino acid units, to form the complex, so proteins and long polypeptides react, while free amino acids and dipeptides do not. The test is named after biuret, a small compound made when urea is heated, which gives the same colour. Sodium hydroxide makes the solution alkaline, and the intensity of the colour is roughly proportional to the number of peptide bonds and so to the amount of protein.

Reagents and Equipment

Name	Specification	Use
Sodium hydroxide solution	About 1 mol/dm³ (4% w/v), corrosive	Makes the solution alkaline
Copper(II) sulphate solution	Dilute, about 1% w/v, pale blue	Supplies copper(II) ions for the complex
Test tubes and rack	Clean, labelled	Hold the mixtures
Dropping pipettes	Separate for each reagent	Add reagents without contamination
Protein solution	1% egg white (albumen) in water, or 1% casein or bovine serum albumin	Positive control
Distilled water	Clean	Negative control

A ready-made Biuret reagent (containing copper sulphate, sodium potassium tartrate and sodium hydroxide) may be used instead; in that case one reagent is added in a single step.

Procedure

1. Put on gloves, goggles and a laboratory coat, because sodium hydroxide is corrosive.
2. Label three tubes: sample, positive control and negative control.
3. Place 2 cm³ of the sample in the first tube, 2 cm³ of protein solution in the second and 2 cm³ of distilled water in the third. A solid food is first ground with water and filtered.
4. Add an equal volume (2 cm³) of sodium hydroxide solution to each tube and mix.
5. Add 2 to 3 drops of dilute copper(II) sulphate solution to each tube, one drop at a time, mixing after each drop.
6. Leave the tubes for about 5 minutes at room temperature. No heating is needed.
7. Observe the colour against a white background and compare it with the controls.
8. Record the colour and the conclusion for each tube.

Results and Interpretation

Colour	Meaning
Blue (the colour of copper sulphate)	No peptide bonds; protein is absent
Violet or purple	Protein or long polypeptide is present
Pink or light purple	Short peptides (peptones, about three to four amino acids) are present

A deeper violet suggests more protein. The test confirms that peptide bonds exist, not which protein it is. Adding too much copper sulphate gives a blue colour that can hide a weak positive, so the drops must be few.

Safety

• Sodium hydroxide is corrosive and burns the skin and eyes, so wear goggles and gloves and rinse splashes with plenty of water at once.
• Copper(II) sulphate is harmful if swallowed and toxic to aquatic life, so keep it out of drains.
• Never pipette by mouth.
• Use separate droppers for each reagent so that bottles are not contaminated.
• Dispose of used mixtures in the labelled waste container.

Sources of Error

• Too much copper sulphate: The strong blue hides the violet colour and gives a false negative.
• Ammonium salts and Tris buffer: These bind copper and also give a violet-like colour, so the sample must not contain them.
• Cloudy samples: Fat globules and suspended solids cloud the solution; milk should be centrifuged or diluted so the colour can be seen.
• Very dilute protein: The test is not sensitive below about 1 mg of protein per cm³, so dilute samples may be missed.
• Reading too soon or too late: The colour needs a few minutes to develop, but standing for a long time can allow copper hydroxide to precipitate.

2.1.5 DCPIP test (ascorbic acid test)

The DCPIP test detects ascorbic acid (vitamin C) and can be used to compare how much of it different foods contain. DCPIP is 2,6-dichlorophenolindophenol, a dye that is blue when oxidised and colourless when reduced. Example: a nutrition researcher at a university can use it to compare the vitamin C in fresh oranges, mangoes and passion fruit juice sold at a market, and to show how much is lost after boiling or storage.

Principle

Ascorbic acid is a strong reducing agent. When it is added to blue DCPIP solution, it donates electrons (it is oxidised to dehydroascorbic acid) and the DCPIP is reduced to a colourless compound. The disappearance of the blue colour is the end point. In acidic solution, such as a fruit juice, the oxidised dye may look pink or red before it turns colourless, and the end point is the loss of that colour. The volume of juice that is needed to decolourise a fixed volume of dye depends on its vitamin C content: the more vitamin C in the juice, the less juice is needed.

Reagents and Equipment

Name	Specification	Use
DCPIP solution	About 0.1% w/v (1 g/dm³) in water, freshly made, deep blue	The indicator dye that is decolourised
Ascorbic acid standard	0.1% w/v (1 mg/cm³), freshly made	Standard of known concentration
Burette or graduated syringe	1 cm³ graduated in 0.01 cm³; or dropper of known drop size	Add juice or standard in measured volumes
Test tubes or small beaker	Clean	Hold the dye
Fruit juices	Freshly squeezed and filtered	Samples
Distilled water	Clean	Negative control and for dilution

Procedure

1. Wear a laboratory coat and gloves, and label the tubes: standard, each juice and water control.
2. Filter each juice through muslin or filter paper to remove pulp, because the particles hide the colour change.
3. Pipette 1.0 cm³ of blue DCPIP solution into a clean tube.
4. Fill the burette or syringe with the ascorbic acid standard and add it to the dye one drop (or 0.1 cm³) at a time, shaking after each addition, until the blue colour just disappears. Record the volume used.
5. Repeat with a fresh 1.0 cm³ of DCPIP for each juice, using the same method and the same size of drops.
6. Put 1.0 cm³ of DCPIP in a tube with distilled water instead of juice as the control; it must stay blue.
7. Repeat each determination three times and calculate the mean volume.
8. Calculate the vitamin C concentration using the standard.

Worked calculation:
Volume of standard (1 mg/cm³) needed for 1.0 cm³ DCPIP = 0.40 cm³
Vitamin C that decolourises 1.0 cm³ DCPIP = 0.40 cm³ × 1 mg/cm³ = 0.40 mg
Volume of orange juice needed for 1.0 cm³ DCPIP = 0.80 cm³
Vitamin C in juice = 0.40 mg ÷ 0.80 cm³ = 0.50 mg/cm³ (50 mg per 100 cm³)

Results and Interpretation

• Blue disappears quickly: A small volume of juice decolourises the dye, which means a high vitamin C content.
• Large volume needed: The juice has a low vitamin C content.
• Blue stays after a large volume: Little or no vitamin C is present, or it has been destroyed by heating or storage.
• Water control stays blue: Shows that the dye itself is stable and that the colour loss in the other tubes is caused by the juice.
• Comparison with the standard: The standard solution gives a reference volume, so the mass of vitamin C in the juice can be calculated.

Safety

• DCPIP stains skin and clothing, so wear gloves and a laboratory coat.
• Avoid contact with the eyes and do not swallow the solution.
• Never taste the juices used in the test, because glassware may be contaminated.
• Wash hands after the practical.
• Dispose of dye solutions as the laboratory directs.

Sources of Error

• Vitamin C breaks down: Ascorbic acid is oxidised by air, heat and light, so juices should be freshly prepared and tested promptly.
• Coloured juice: Dark juices hide the end point; dilute the juice or use a colourless extract.
• Other reducing substances: Sulphites, thiols and some other compounds also decolourise DCPIP, so the result is the reducing capacity, not only vitamin C.
• Different drop sizes: Drops from different droppers are not the same volume, so a burette or one syringe should be used.
• Old DCPIP solution: The dye fades on standing, so it should be made fresh and kept in the dark.

2.1.6 Chromatography

Chromatography is a family of techniques that separate the components of a mixture as they are carried by a mobile phase through or over a stationary phase. Each component moves at its own speed, so the mixture spreads into separate zones. Example: a public analyst at a university can separate the pigments in a leaf extract on paper, an agricultural testing laboratory can separate food dyes in juice, and a research laboratory can analyse amino acids from a protein hydrolysate.

Principle

Substances in a mixture are distributed between two phases. The stationary phase is a solid or a liquid held on a solid (the paper, the silica or the packing), and the mobile phase is the liquid or gas that moves through it. A substance that dissolves well in the mobile phase and is held weakly by the stationary phase travels far. A substance that is held strongly by the stationary phase travels only a short way. In paper chromatography, the stationary phase is the water held in the cellulose fibres of the paper, and separation is mainly by partition between that water and the solvent. In thin layer chromatography, the stationary phase is a thin layer of silica or alumina on a plate, and separation is mainly by adsorption.

Reagents and Equipment

Name	Specification	Use
Chromatography paper	Whatman No. 1 or equivalent strips, about 15 cm × 3 cm	Stationary phase
Developing tank	Glass jar or beaker with a lid or cling film	Keeps solvent vapour saturated
Solvent (mobile phase)	For pigments: petroleum ether and propanone, about 9:1; for amino acids: butan-1-ol, ethanoic acid and water, 4:1:5	Carries the substances up the paper
Capillary tube or fine pipette	Fine glass	Applies small spots
Pencil and ruler	Soft pencil, mm ruler	Mark the origin and the front, measure distances
Locating agent	Ninhydrin solution, about 0.2% in propanone	Makes colourless amino acids visible
Leaf extract or sample	Crushed in propanone with sand	Sample for separation

Procedure

1. Wear gloves, goggles and a laboratory coat, and work in a well-ventilated area away from flames.
2. Handle the paper only by the edges, because fingerprints contain amino acids and oils.
3. Draw a faint pencil line (the origin) about 2 cm from the bottom of the strip, and mark the positions of the spots on it. Do not use ink, which would run in the solvent.
4. Crush fresh leaves with a little propanone and sand and filter, to obtain a concentrated extract.
5. Use a capillary tube to place a small spot on the origin. Let it dry and apply again 5 to 10 times in the same place to build a concentrated, small spot.
6. Pour solvent into the tank to a depth of about 1 cm, so that the level is below the pencil line. Cover the tank and wait a few minutes for the vapour to saturate.
7. Hang the strip so that the bottom dips into the solvent without the spots touching it, and replace the lid.
8. Remove the strip when the solvent front is about 1 to 2 cm from the top, and mark the position of the front with a pencil at once, before it dries.
9. Dry the strip. If the substances are colourless (such as amino acids), spray with ninhydrin and warm it in an oven at about 100 °C for a few minutes to show purple spots (proline gives yellow).
10. Mark the centre of each spot with a pencil and measure the distances.

Rf Value and Worked Example

The retention factor (Rf) compares how far a substance has moved with how far the solvent has moved. It has no unit and is always between 0 and 1.

Rf = distance moved by the substance from the origin ÷ distance moved by the solvent front from the origin

Worked example: a leaf extract is run in petroleum ether and propanone. The solvent front moves 10.0 cm from the origin. The spots are measured from the origin to the centre of each spot.

Carotene: distance 9.5 cm, Rf = 9.5 ÷ 10.0 = 0.95
Xanthophyll: distance 7.5 cm, Rf = 7.5 ÷ 10.0 = 0.75
Chlorophyll a: distance 6.0 cm, Rf = 6.0 ÷ 10.0 = 0.60
Chlorophyll b: distance 4.5 cm, Rf = 4.5 ÷ 10.0 = 0.45

Carotene has the highest Rf because it is the least polar pigment and dissolves best in the non-polar solvent, while chlorophyll b is the most polar and is held most strongly by the paper. The exact values depend on the solvent, the paper and the temperature, so substances are compared with standards run on the same strip under the same conditions. A second example: a spot that moves 3.2 cm when the solvent front moves 8.0 cm has Rf = 3.2 ÷ 8.0 = 0.40.

Other Types of Chromatography

• Thin layer chromatography (TLC): A silica plate is used instead of paper, giving faster and sharper separations and used for dyes, drugs and lipids.
• Column chromatography: The mixture is passed through a column packed with silica or resin, and fractions are collected as they emerge; it is used to purify larger amounts.
• Ion-exchange chromatography: Charged resin beads hold molecules of opposite charge, which are then released with a salt gradient; it is used to separate proteins and amino acids.
• Size exclusion (gel filtration) chromatography: Porous beads let small molecules enter and so delay them, while large molecules pass quickly through; it separates molecules by size.
• High performance liquid chromatography (HPLC) and gas chromatography (GC): A pump forces a liquid, or a carrier gas carries a vapour, through a packed column, with a detector recording each component; they are used for vitamins, pesticide residues and drugs.

Safety

• Propanone, petroleum ether and butan-1-ol are flammable and their vapours are harmful, so work in a ventilated area away from flames and sparks.
• Wear gloves and goggles, because the solvents irritate the skin and eyes.
• Ninhydrin is an irritant and stains the skin; spray in a fume cupboard or a well-ventilated area and avoid breathing the mist.
• Keep the developing tank covered so that vapour does not build up in the room.
• Dispose of the solvents in a labelled waste container and never pour them into drains.

Sources of Error

• Origin line under the solvent: If the starting spots are below the solvent level, the sample dissolves into the tank instead of moving up the paper.
• Spots too large or too dilute: Large spots overlap and diffuse, so the separation is poor and the Rf is hard to measure.
• Unsaturated tank: The solvent evaporates from the paper and the front runs unevenly, changing the Rf.
• Failing to mark the solvent front: Once the strip dries, the front cannot be seen and the Rf cannot be calculated.
• Touching the paper with fingers or letting it touch the sides: Contamination and uneven flow give false spots and crooked fronts.

2.1.7 Electrophoresis

Electrophoresis separates charged molecules, such as DNA, RNA and proteins, by their movement in an electric field through a gel. It is used to check PCR products, to separate DNA fragments cut by restriction enzymes, to analyse proteins in blood serum and to identify genetic variants. Example: a research institute that screens maize varieties for disease resistance genes uses agarose gel electrophoresis to see whether a PCR product of the expected size is present.

Principle

Molecules with a net charge move in an electric field: negatively charged molecules move towards the positive electrode (anode) and positively charged molecules towards the negative electrode (cathode). The gel is a porous network that acts like a sieve, so small molecules move through quickly and large molecules are slowed. DNA has a negative charge because of its phosphate groups, and because the charge per unit length is the same for every fragment, the fragments are separated by size: the smallest travel furthest. For proteins, sodium dodecyl sulphate (SDS) is added to coat each protein with negative charge in proportion to its mass, so SDS polyacrylamide gel electrophoresis (SDS-PAGE) also separates by size and not by the protein's own charge. The rate of movement depends on the size of the molecule, its charge, the applied voltage, the gel concentration and the buffer.

Gel Types

• Agarose gel: Made from a seaweed polysaccharide and used for DNA and RNA; large pores suit fragments from about 100 base pairs to over 20,000. A 0.7% gel separates large fragments, 1% is a general-purpose gel and 2% separates small fragments.
• Polyacrylamide gel: Made by polymerising acrylamide and bis-acrylamide, giving smaller and more uniform pores; used for proteins (SDS-PAGE) and for very small DNA fragments. A 12% gel suits proteins of about 15 to 60 kDa.
• Native gels: Run without SDS, so proteins keep their folded shape and activity and move according to their own charge, size and shape.
• Denaturing gels: Contain SDS (for proteins) or urea (for nucleic acids) so that molecules are unfolded and separated by length or mass only.
• Stacking and resolving layers: SDS-PAGE uses a low-percentage stacking gel at about pH 6.8 to line the proteins up into a thin band, and a higher-percentage resolving gel at about pH 8.8 to separate them.

Reagents and Equipment

Name	Specification	Use
Agarose	Molecular biology grade, 0.7 to 2% w/v	Forms the gel
Electrophoresis buffer	TAE (40 mmol/dm³ Tris-acetate, 1 mmol/dm³ EDTA, about pH 8.3) or TBE	Carries the current and keeps pH steady
Gel tank, tray and comb	Horizontal tank, casting tray, comb for wells	Holds and shapes the gel
Power supply	Direct current, about 80 to 120 V	Provides the electric field
Loading dye	Glycerol or sucrose with bromophenol blue	Makes samples sink in the wells and shows progress
DNA marker (ladder)	Fragments of known sizes	Used to estimate sizes
Stain	Ethidium bromide (about 0.5 µg/cm³) or a safer dye such as SYBR Safe	Makes DNA visible
UV transilluminator or gel imager	Ultraviolet light, about 300 nm	Shows stained bands
Micropipette	2 to 20 µL	Loads the samples

Procedure

1. Put on a laboratory coat, gloves and goggles, because the stain and the voltage are hazardous.
2. Weigh the agarose (for example 1.0 g for a 1% gel in 100 cm³ of buffer), add the buffer in a flask, heat in a microwave or hot water bath until the solution is clear, and cool to about 55 °C.
3. Add the stain to the cooled agarose (if used in the gel), mix gently, and pour into a tray with the comb in place so that wells form. Let it set for about 30 minutes.
4. Remove the comb and place the set gel in the tank. Cover it with buffer until it is about 3 to 5 mm above the gel.
5. Mix each sample with loading dye (for example 5 µL of sample and 1 µL of dye).
6. Load the samples into the wells with a micropipette, and load the marker (ladder) into one well. Record the order of the samples.
7. Close the lid and connect the electrodes so that the wells are at the cathode (negative) end. The DNA moves towards the anode (positive).
8. Run at about 5 V per cm between the electrodes (typically 80 to 100 V) for 30 to 60 minutes, until the loading dye has moved about two thirds of the gel.
9. Switch off the power, remove the gel, and stain it if the stain was not in the gel. For SDS-PAGE, stain with Coomassie Brilliant Blue, then destain with methanol and ethanoic acid solution until the bands are clear.
10. View under UV light (for ethidium bromide or similar dyes) or on a white background (for Coomassie), and photograph the gel.

Interpretation of Results

• Band position: A band that is further from the well is a smaller fragment; a band near the well is a large one.
• Comparison with the marker: The sizes of the ladder fragments are known, so each sample band can be matched to a size by comparing distances moved. A plot of the log of the fragment size against the distance moved gives a straight line that is used to read the size of an unknown.
• Band intensity: A thicker or brighter band suggests more DNA, but this is only a rough guide to the amount.
• Smear: A smear instead of a sharp band suggests degraded DNA or an overloaded well.
• Absence of a band: The target may be missing, the reaction failed, or too little sample was loaded.
• Single band in a PCR lane: Suggests one product of the expected size; extra bands show non-specific products.

Safety

• Ethidium bromide is a mutagen, so wear gloves, handle with care and dispose of it as hazardous waste; safer dyes should be used where available.
• UV light damages the eyes and skin, so wear a face shield or view through a protective screen.
• The power supply is a risk of electric shock: keep the lid on, switch off before touching the tank, and keep liquids away from the connections.
• Hot agarose can scald, and the flask should be swirled with a protective glove.
• Acrylamide is a neurotoxin, so unpolymerised acrylamide is handled only with gloves and in ventilated areas.

Sources of Error

• Wrong gel concentration: A low percentage gel cannot resolve small fragments, and a high percentage gel slows large fragments too much.
• Wrong buffer or old buffer: Exhausted buffer changes the pH and the current, causing smeared or distorted bands.
• Overloading the wells: Too much sample causes smears and bands that run together.
• Excess voltage: The gel overheats and bands become curved ("smiling") or smeared, and the gel may melt.
• Uneven wells or air bubbles in the gel: These distort the lanes and make comparisons unreliable.
• Running the dye off the gel: Small fragments run out of the gel and are lost.

Chapter Summary

This chapter covered how biomolecules are separated and identified in the laboratory. It began with the separation procedures that prepare a sample: breaking open cells in a buffer (cell lysis) and separating cell parts by their size and density with the centrifuge and the ultracentrifuge. It then covered the qualitative tests that identify the main groups of biomolecules: the iodine test for starch, Benedict's test for reducing sugars, the translucent (grease spot) test for lipids, the Biuret test for proteins and the DCPIP test for ascorbic acid. Finally it covered two separation techniques that split a mixture into its parts: chromatography, which separates substances by how they distribute between a stationary and a mobile phase, and electrophoresis, which separates charged molecules by their size and charge in an electric field. A technologist who masters these steps can prepare samples safely, choose the right test or technique, record results correctly and report them clearly in a food, clinical, agricultural or research laboratory.

Am I competent?

At the start of this chapter we promised you would be able to:

• Assemble biomolecule test samples correctly for separation and qualitative analysis.
• Prepare biomolecule collection tools properly according to laboratory procedures.
• Conduct biomolecule separation procedures safely and accurately.
• Record and report biomolecule separation results clearly and precisely.

Tick each one you can genuinely do.

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