bioanalytical chemistry (chem311)- lab manual

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Department of Chemistry

Faculty of Science

University of Hail, Hail, KSA

BIOANALYTICAL CHEMISTRY

CHEM 331 Lab Manual

PREPARED BY

Dr. Saravanan Rajendrasozhan, Assistant Professor of Biochemistry, Department of Chemistry, University of Hail, Hail, Saudi Arabia.

OBJECTIVE

The course aims to give students insight into and an understanding of the analytical chemical problems and procedures related to biological sample analysis. Students should gain an understanding of the principles of advanced analytical techniques as well as the advantages and disadvantages of the individual methods.

COURSE MATERIAL

1. Principles and reactions of protein extraction, purification, and characterization By Hafiz Ahmed. CRC Press, 2005. ISBN: 9780849320347.

2. Understanding Bioanalytical Chemistry: Principles and Applications By Victor Gault and Neville McClenaghan. Wiley annotated edition, 2009. ISBN: 0470029064.

INDEX

ExperimentNumber

EXPERIMENT PAGE

- Lab safety and Introduction 3

1 Qualitative tests for carbohydrates 6

2 Qualitative tests for proteins 9

3 Qualitative tests for lipids 12

4 Determination of protein concentration by Biuret method 15

5 Determination of protein concentration by Coomassie Blue (Bradford) method

17

6 Estimation of DNA concentration by Diphenylamine method 19

7 Determination of carbohydrates by phenol–sulfuric acid method 21

8 Estimation of carbohydrates by dinitrosalicylic acid method 23

9 Separation and identification of amino acids by paper chromatography 24

10 Separation and identification of DNA by agarose gel electrophoresis 26

Bioanalytical Chemistry – Lab manual Page: 2

LABORATORY SAFETY RULES

Laboratories are interesting and potentially dangerous place to work. Laboratory Safety is a very important aspect of science. Without it, experimentation could result in very serious injury. To reduce the risks involved with experimentation, there are certain procedures that we should all follow.

“Safety is the first priority in our lab”

1. Behave responsibly. The dangers of spilled acids, chemicals and broken glassware created by thoughtless actions are too great to be tolerated.

2. Wear lab coat. A lab coat should be worn during laboratory experiments. It provides protection at all times.

3. Dress properly. Long hair and loose clothing (such as ties and ghutra) are a hazard in the laboratory. They may either catch fire or be chemically contaminated. Long hair must be tied back, baggy clothing must be secured.

4. Wear shoes. Shoes must completely cover the foot. Sandals are prohibited because of the hazard from the chemical spills.

5. Wear eye protector. Wear appropriate eye protection (goggles) at all times in the laboratory and in any area where chemicals are stored or handled. Such protection will protect you against chemical splashes.

6. Do not wear contact lenses (even with safety goggles). Contact lenses prevent rinsing chemical splashes from the eye. Vapors in the laboratory (eg. HCl) dissolve in the liquids covering the eye and concentrate behind the lenses. ‘Soft’ lenses are especially bad as chemicals dissolve in the lenses themselves and are released over several hours.

7. Do not smoke. Smoking is not just an obvious fire hazard; it also draws chemicals in laboratory air (both vapors and dust) into the lungs.

8. Do not eat or drink. Do not eat food, drink beverages, or chew gum in the laboratory. Do not use laboratory glassware as containers for food or beverages. Wash your hands thoroughly with soap and water when leaving the laboratory.

9. Avoid breathing fumes of any kind. To test the smell of a vapor, collect some in a cupped hand. Obtain your instructor’s written permission before you smell any chemical. Never smell a chemical reaction while it is occurring. Work in a hood if there is the possibility that noxious vapors may be produced.


10. Be prepared for your work in the laboratory. Carefully read the experiment before coming to the laboratory. An unprepared student is a hazard to everyone in the room.

11. KEEP YOUR WORK AREA NEAT AND CLEAN. Clean up spills and broken glass immediately. Clean up your work space; wipe all surfaces and put away all chemicals and equipment at the end of the laboratory period.

12. Never work alone in the lab. There must be at least one other person present in the lab. In addition, an instructor should be quickly available.

13. Do not perform any unauthorized experiments. Perform only those experiments authorized by your teacher. This includes using only the quantities instructed, no more. Carefully follow all instructions, both written and oral. Consult your instructor if you have any doubts about instructions and laboratory manuals.

14. Be careful while heating liquids. Add boiling chips to avoid ‘bumping’. Flammable liquids such as ethers, hydrocarbons, alcohols, acetone, and carbon disulfide must never be heated over direct flame.

15. Careful about spatters. Spatters are common in general chemistry lab. Test tubes being heated or containing reaction mixtures should never be pointed at anyone. If you observe this practice in a neighbor, speak to him or the instructor.

16. Always pour acids into water when mixing. Otherwise the acid can spatter, often quite violently.

17. Never use mouth suction in filling pipettes with chemical reagents. Always use a suction device.

18. Do not force a rubber stopper onto glass tubing or thermometers. Lubricate the tubing and the stopper with glycerol or water. Use paper or cloth toweling to protect your hands. Grasp the glass close to the stopper.

19. Dispose of all chemical waste properly. Dispose of excess liquid reagents by flushing small quantities down the sink. Consult the instructor about large quantities. Dispose of solid in crocks. Never return unused chemicals to their original container.

20. Report any accident (spill, breakage, etc.) or injury (cut, burn, etc.) to your instructor immediately. Do not panic.

21. In case of fire or accident, call the instructor at once. Note the location of fire extinguisher and safety showers now so that you can use them if needed.

- Wet towels can be used to smother small fires.


- In case of chemical spill on your body or clothing, wash the affected area with large quantities of running water. Remove clothing that has been wet by chemicals to prevent further reaction with the skin.

- If a chemical should splash in your eye(s), immediately flush with running water for at least 20 minutes.

- Except for superficial injuries, you will be required to get medical treatment for cuts, burns or fume inhalation.

22. Most importantly, think about what you are doing. Plan ahead. If you give no thought to what you are doing, you predispose yourself to an accident.

23. Report if you have any diagnosed allergies or other special medical needs to your instructor.

INTRODUCTION

Bioanalytical chemistry

A branch of analytical chemistry in which compounds of biological significance, such as

carbohydrates, proteins, lipids and nucleic acids, are studied.

Qualitative analysis

It deals with the identification of elements or grouping of elements (example: Biomolecules)

present in a sample.

Quantitative analysis

It deals with the determination of the absolute or relative abundance (often expressed as a

concentration) of a particular substance present in a sample.


Experiment: 1

QUALITATIVE ANALYSIS OF CARBOHYDRATES

1. BENEDICT’S TEST

Principle

Glucose reduces copper hydroxide (cupric hydroxide) in alkaline solution to red or brown colored cuprous oxide. The reducing property of glucose is due to the presence of free aldehyde (or keto) group. Depending on the concentration, yellow to green color is developed.

Reagents

1. Benedict’s reagent

2. Test solution: 10 grams of glucose in 1000 ml of distilled water.

Procedure

Add 5 drops of the test solution to 2 ml of Benedict’s reagent. The solution is boiled for 5 minutes in a water both. Cool the solution.

Analysis

Formation of red, yellow or green precipitate indicates the presence of glucose.

2. FEHLING’S TEST

Principle

When the blue alkaline cupric hydroxide present in Fehling’s solution is heated with glucose, it gets reduced to yellow or red cuprous oxide precipitate.

Reagents

1. Fehling’s solution A and B

2. Test solution: 10 grams of glucose in 1000 ml of distilled water.

Procedure

To 1 ml of Fehling’s solution A, add 1 ml of Fehling’s solution B and a few drops of test solution. Boil for a few minutes.

Analysis

Formation of brownish-red precipitate indicates the presence of glucose.


3. IODINE TEST

Principle

When starch solution is mixed with iodine, a blue color starch-iodine complex is formed.

Reagents

1. Iodine solution: 2 g of potassium iodide is dissolved in 100 ml of distilled water. To this 62 mg of iodine crystals are dissolved (or 3% potassium iodide).

2. Test solution: 1 gm of starch powder is boiled in 100 ml of distilled water.

Procedure

Take 2 ml of starch solution in a test tube. To these 3 drops of iodine solution is added.

Analysis

Formation is deep blue color indicates the presence of starch.

4. ANTHRONE TEST

Principle

Carbohydrates are first hydrolyzed into sample sugars using dilute HCl. In hot acidic medium glucose is dehydrated to hydroxymethylfurfural. This compound with anthrone forms a green colored product with an absorption maximum at 630 nm.

Reagents

1. 2.5 N Hydrochloric acid.

2. Anthrone reagent: 200 mg of anthrone is dissolved in 100 ml of concentrated sulfuric acid.

3. Test solution: Dissolve 1 gm table sugar or glucose or starch in 100 ml of distilled water.

Procedure

Take about 2 ml of test solution in a test tube. To this 2 ml of anthrone reagent is added and mixed well. Heat for 8 min in a water bath. Cool the test tube.

Analysis

Formation is dark green color indicates the presence of carbohydrates.


Date:

REPORT

Experiment 1: QUALITATIVE ANALYSIS OF CARBOHYDRATES

Test Observation Conclusion

1. BENEDICT’S TEST

2. FEHLING’S TEST

3. IODINE TEST

4. ANTHRONE TEST

Result

The given sample contains __________________________________________________________


Experiment: 2

QUALITATIVE ANALYSIS OF PROTEINS

1. NINHYDRIN TEST

Principle

Ninhydrin is a powerful oxidizing agent. It decarboxylate the alpha amino acids and yields an intensely colored bluish purple product.

Reagents

1. 1% Ninhydrin: Dissolve 1 g of ninhydrin in 100 ml of acetone.

2. Test solution: Dissolve 1 g of peptone in 100 ml of distilled water (1% solution).

Procedure

Take 2 ml of test solution is a test tube. To this add 1 ml of 1% freshly prepared ninhydrin solution. Mix the contents and boil for a couple of minutes. Allow to cool.

Analysis

Formation of violet or purple color indicates the presence of protein.

2. BIURET TEST

Principle

Compounds with two or more peptide bonds give a violet color with alkaline copper sulphate solution.

Reagents

1. Biuret reagent

2. Test solution: Dissolve 1 g of peptone in 100 ml of distilled water (1% solution) or separate the egg white and dilute it.

Procedure

Take 2 ml of test solution. To this add 2 ml of biuret reagent.

Analysis

Formation of violet or pink color indicates the presence of protein.Bioanalytical Chemistry – Lab manual Page: 9

3. NITRIC ACID TEST

Reagents

1. Nitric acid

3. Test solution: Separate the egg white and dilute it.

Procedure

Take 2 ml of test solution in a test tube. Add a few drops of nitric acid. A precipitate appears. Heat it.

Analysis

Formation of lemon yellow color indicates the presence of protein.


Date:

REPORT

Experiment 2: QUALITATIVE ANALYSIS OF PROTEINS


1. NINHYDRIN TEST

2. BIURET TEST

3. NITRIC ACID TEST

Result



Experiment: 3

QUALITATIVE ANALYSIS OF LIPIDS

1. SOLUBILITY TEST

Principle

Lipids dissolve in chloroform, not in water.

Reagents

1. Chloroform

2. Test solution: Coconut oil or ghee

Procedure

Take 2 drops of the sample in two separate test tubes. Add 2 ml of distilled water to one tube and 2 ml of chloroform to another tube. Shake well and observe.

Analysis

If the sample completely dissolved in chloroform and undissolved in water, it indicates the presence of lipid.

2. EMULSIFICATION TEST

Reagents

1. 5% Sodium carbonate


Procedure

Take 2 drops of the sample and 4 ml of distilled water in a test tube. Add 4 drops of 5% sodium carbonate. Shake well and observe.

Analysis

Formation of emulsion (uniform distribution of lipid in solution) indicates the presence of lipid.


3. IODINE TEST

Reagents

1. Iodine solution: Dissolve 2 g of potassium iodide in 100 ml of distilled water. To this add 62 mg of iodine crystals and dissolve.


Procedure

Take 5 drops of test solution in a test tube. Add 1 ml of distilled water and 1 ml of iodine. Shake well.

Analysis

Disappearance of yellow color (due to the absorption of iodine by lipids) indicates the presence of lipid.

4. SUDAN III TEST

Reagents

1. Sudan III.


Procedure

Take 2 ml of test solution and add few drops of sudan III.

Analysis

Formation of red color indicates the presence of lipid.


Date:

REPORT

Experiment 3: QUALITATIVE ANALYSIS OF LIPIDS


1. SOLUBILITY TEST

2. EMULSIFICATION TEST

3. IODINE TEST

4. SUDAN III TEST

Result



Experiment: 4

DETERMINATION OF PROTEIN CONCENTRATION BY BIURET METHOD

Principle

Under alkaline conditions substances containing two or more peptide bonds (proteins) form a purple complex with copper salts in the reagent.

Equipment

In addition to standard liquid handling supplies a visible light spectrophotometer is needed, with maximum transmission in the region of 450 nm. Glass or polystyrene cuvettes may be used.

Reagent

1. Biuret reagent: 9 gm Sodium potassium tartrate (f.w. 282.22), 3 gm Copper sulfate x 5 H2O (f.w. 249.68), 5 gm Potassium iodide (f.w.166.0), all dissolved in order in 400 ml 0.2 M NaOH (f.w. 40.0) before bringing to final volume (one liter final volume). Discard if a black precipitate forms.

Assay Procedure

1. Warm up the spectrophotometer 15 minutes before use. 2. Prepare standards from bovine serum albumin (recommended range is 0.5 to 20 mg protein).3. Prepare a reference tube with 1 ml buffer.4. If possible, dilute unknowns to an estimated 1 to 10 mg/ml with buffer; a range of dilutions

should be used if the actual concentration cannot be estimated.5. Use 1 ml sample per assay tube.6. Add 9 ml Biuret reagent to each tube, vortex immediately, and let stand 20 min.7. Read at 550 nm.


Analysis

Prepare a standard curve of absorbance versus micrograms protein (or vice versa). From the curve, determine concentrations of unknown samples from the amount protein, volume/sample, and dilution factor, if any.


Experiment: 5

DETERMINATION OF PROTEIN CONCENTRATION BY COOMASSIE BLUE (BRADFORD) METHOD

Principle

The assay is based on the observation that the absorbance maximum for an acidic solution of Coomassie Brilliant Blue G-250 shifts from 465 nm to 595 nm when binding to protein occurs. Both hydrophobic and ionic interactions stabilize the anionic form of the dye, causing a visible color change. The assay is useful since the extinction coefficient of a dye-albumin complex solution is constant over a 10-fold concentration range.

Equipments

In addition to standard liquid handling supplies a visible light spectrophotometer is needed, with maximum transmission in the region of 595 nm, on the border of the visible spectrum (no special lamp or filter usually needed). Glass or polystyrene cuvettes may be used, however the color reagent stains both. Disposable cuvettes are recommended.

Reagents

1. Bradford reagent: Dissolve 100 mg Coomassie Brilliant Blue G-250 in 50 ml 95% ethanol, add 100 ml 85% (w/v) phosphoric acid. Dilute to 1 liter when the dye has completely dissolved, and filter through Whatman #1 paper just before use.

2. (Optional) 1 M NaOH (to be used if samples are not readily soluble in the color reagent).


The Bradford reagent should be a light brown in color. Filtration may have to be repeated to rid the reagent of blue components. The Bio-Rad concentrate is expensive, but the lots of dye used have apparently been screened for maximum effectiveness. "Homemade" reagent works quite well but is usually not as sensitive as the Bio-Rad product.

Assay procedure

1. Warm up the spectrophotometer before use.2. Dilute unknowns if necessary to obtain between 5 and 100 µg protein in at least one assay

tube containing 100 µl sample.3. Add an equal volume of 1 M NaOH to each sample and vortex. 4. Prepare standards containing a range of 5 to 100 micrograms protein (albumin or gamma

globulin are recommended) in 100 µl volume. 5. Add 5 ml dye reagent and incubate 5 min.6. Measure the absorbance at 595 nm.

Analysis

Prepare a standard curve of absorbance versus micrograms protein and determine amounts from the curve. Determine concentrations of original samples from the amount protein, volume/sample, and dilution factor, if any.


Experiment: 6

ESTIMATION OF DNA CONCENTRATION BY DIPHENYLAMINE METHOD

Principle

The reagents used in the diphenylamine reaction include acetic acid and sulfuric acid. When these are heated with DNA they cleave the phospho-diester bonds and hydrolyze the glycosidic bonds between the sugar and purines. The free 2-deoxy ribose undergoes a dehydration reaction to form ω-hydroxylevulinyl aldehyde, which reacts with diphenylamine to produce a variety of blue-colored compounds showing a characteristic absorbance peak at 595 nm.

Since the reaction is specific for 2-deoxy ribose, the sugar in DNA, there is no reaction with the ribose sugar of RNA. Thus, the presence of RNA in a sample will not interfere with the measurement of DNA.

Equipments

In addition to standard liquid handling supplies, boiling water bath and visible light spectrophotometer are needed, with maximum transmission in the region of 595 nm, on the border of the visible spectrum (no special lamp or filter usually needed).

Reagents

1. Diphenylamine reagent: Dissolve 1 g of diphenylamine in 100 ml of glacial acetic acid, and subsequently add 2.5 ml conc. H2SO4. Always use freshly prepared reagent and cover it with an aluminium foil, as the reagent is photosensitive.

2. Stock solution: Prepare a stock solution with 100 g/ml of commercially available calf thymus DNA in distilled water or buffered saline.

Assay procedure

1. Prepare a series of standard DNA dilutions of 0 to 100 g in 1 ml of each using stock solution.2. Set up various dilutions of unknown DNA.3. Add 3 ml of diphenylamine reagent to all test tubes containing the diluted DNA samples and

blank.

Note: diphenylamine reagent contains sulfuric and acetic acids. Use extreme caution when handling. Clean up spills with plenty of water.

4. Mix the contents, and cap the tubes with foil, wrap with tape, and label each tube using a pencil. Place the tubes in a boiling water bath for 10 minutes (do not submerge tubes in water!).


5. Remove the tubes from the boiling water and cool them briefly in an ice bath, then allow them to reach room temperature.

6. Once the tubes are dry, read their absorbance at 600 nm.

Analysis

Prepare a standard curve of absorbance versus micrograms DNA and determine amounts from the curve. Determine concentrations of original samples from the amount of DNA, volume/sample, and dilution factor, if any.


Experiment: 7

DETERMINATION OF CARBOHYDRATES BY PHENOL–SULFURIC ACID METHOD

Among many colorimetric methods for carbohydrate analysis, the phenol–sulfuric acid method is the easiest and most reliable method. It has been used for measuring neutral sugars in oligosaccharides, proteoglycans, glycoproteins, and glycolipids. This method is used widely because of its sensitivity and simplicity.

Principle

This assay based on the action of concentrated sulfuric acid that causes hydrolysis of glycosidic linkage. The hydrolysed neutral sugar (pentoses and hexoses) are then partially dehydrated, with the elimination of three molecules of water, to form furfural or a derivative of furfural (hydroxyl methyl furfural).

(Hexose)

Furfural or hydroxyl methyl furfural cause condensation with phenol to form yellow-orange colored compounds. The absorbance at 420 nm is proportional to the carbohydrate concentration initially in the sample. The sulfuric acid causes all non-reducing sugars to be converted to reducing sugars, so that this method determines the total sugars present.

Reagents

1. Phenol solution: 5% w/v in water.

Assay procedure

1. Set up several glass tubes containing 5 to 100 g glucose in 200 l of water. Take unknown glucose in 200 l of water.

2. Add 200 l of phenol solution.3. Add 1 ml concentrated sulfuric acid rapidly and directly on the sample (Do not allow sulfuric

acid to touch the side of the tube).


4. Leave the solution undisturbed for 10 minutes.5. Shake vigorously and continue incubation for another 30 minutes.6. Read each tube at 490 nm.

Analysis

Prepare a standard curve of absorbance versus micrograms carbohydrate (glucose). Determine the concentration of carbohydrate in the unknown sample from a standard plot.


Experiment: 8

ESTIMATION OF CARBOHYDRATES BY DINITROSALICYLIC ACID METHOD

Principle

3,5-Dinitrosalicylic acid (DNS) is an aromatic compound that reacts with reducing sugars and other reducing molecules to form 3-amino-5-nitrosalicylic acid, which absorbs light strongly at 540 nm.

It was first introduced as a method to detect reducing substances in urine and has since been widely used, for example, for quantificating carbohydrates levels in blood.

Reagents

1. Assay reagent: Dissolve 0.25 g 3,5-dinitrosolicylic acid and 75 g sodium potassium tartrate in 50 ml 2M NaOH. Dilute to 250 ml with water.

Assay procedure

1. Set up several glass tubes containing 5 to 500 g glucose in 100 l water. Set up tube for unknown sample in 100 l water.

2. Add 1 ml of assay reagent in to each tube. Mix well.3. Incubate in boiling water bath for 10 minutes.4. Cool to room temperature and determine absorbance at 570 nm.

Analysis

Prepare a standard curve of absorbance versus micrograms carbohydrate (glucose). Determine the concentration of carbohydrate in the unknown sample from a standard plot.


Experiment: 9

SEPARATION AND IDENTIFICATION OF AMINO ACIDS BY PAPER CHROMATOGRAPHY

Principle

Chromatography is a technique for separating mixtures into the components that they are made from. Chromatography utilizes the differential affinities of the components for a gas or liquid mobile medium (mobile phase) and for a stationary adsorbing medium (stationary phase) through which they pass. The stationary phase holds the mixture until the mobile phase passes through, solubilizes the components, and moves them along at their individual rates. Once components are separated from one another, they can be analyzed.

In paper chromatography the stationary phase is the filter paper and the mobile phase is the solvent. The filter paper holds the components until the solvent dissolves them and carries them up the filter paper. The solvent travels up the filter paper by capillary action. The solvent’s attraction to itself pulling it up is greater than the force of gravity pulling it down. The separation of components depends on their solubility with the solvent and their affinity to the solvent and filter paper.

Objectives

1. To demonstrate separation of amino acids by paper chromatography.2. To use chromatography to identify amino acids.

Materials

1. Chromatography tank and lid2. Chromatography paper (or filter paper), to fit tank3. Capillary tubes or pasteur pipettes4. Amino acid samples (e.g. Leucine and Glycine) in 1 molar hydrochloric acid5. Solvent (BAW): Butan-1-ol : Acetic acid : Water 60 : 15 : 25 (should be made up fresh on the

day)6. Spray can of 1% Ninhydrin in butan-1-ol7. Latex gloves

Experimental Procedure

1. Using a pencil, lightly draw a line 1.5–2 cm above the bottom of the chromatography paper. Make small marks at 1.5 cm intervals along the line.

2. Fill a capillary tube or pipette by capillary action with your sample (animo acid). Touch the tip of the capillary to the first mark and pull it away. You should leave a small (>5 mm) wet


spot. Let this dry and apply the sample again. Make a note of which sample is placed on the spot (you can lightly write on the paper beneath the mark, using a pencil). Repeat for each sample or mixture, on a different mark and using a fresh tube or pipette each time. Let the paper dry.

3. Put 1 cm or less of BAW into the chromatography tank. The sample spots should not dip into the solvent. Place the tank in a cool place out of direct sunlight. Stand the chromatography paper in the tank so that the bottom edge is in the solvent but the remaining paper does not touch the tank. Place the lid on the tank and leave until the solvent reach the top.

4. Wearing gloves, remove the damp paper and mark where the solvent has reached. Dry the paper.

5. Spray the whole of the paper with 1% ninhydrin. Dry the paper.6. Mark the position of each spot that develops.

Analysis

Note the color and travel distance of each standard and hence find the amino acid composition of the unknown samples.

The relative extent to which solute molecules (amino acids) move in a chromatography experiment is indicated by retardation factor (Rf) values. Calculation of Rf of the amino acids:

Rf = Distance travelled by amino acid (a or b) / Distance travelled by solvent (s)


Experiment: 10

SEPARATION AND IDENTIFICATION OF DNA BY AGAROSE GEL ELECTROPHORESIS

Agarose gel electrophoresis is a widely used technique for the analysis of nucleic acids. DNA agarose gels can be used to separate and visualize DNA fragments of various sizes.

Principle

The technique of DNA agarose gel electrophoresis is based on the fact that DNA is negatively charged at neutral pH due to its phosphate backbone. For this reason, when an electrical potential is placed on the DNA it will move toward the positive pole.

An agarose gel is created by suspending dry agarose in a buffer solution, boiling until the solution becomes clear, and then pouring it into a casting tray and allowing it to cool. The result is a flexible gelatin-like slab. During electrophoresis, the gel is submersed in a chamber containing a buffer solution and a positive and negative electrode. The DNA to be analyzed is forced through the pores of the gel by the electrical current. Under an electrical field, DNA will move to the positive electrode (red) and away from the negative electrode (black). Several factors influence how fast the DNA moves, including; the strength of the electrical field, the concentration of agarose in the gel and most importantly, the size of the DNA molecules. Smaller DNA molecules move through the agarose faster than larger molecules. DNA in the gel will be visualized by the use of Ethidium Bromide, added to the gel. Ethidium bromide binds to DNA and illuminates when exposed to ultraviolet light, causing the DNA to ‘glow’.

Materials

1. Agarose2. TAE Buffer: Add 4.84 g Tris Base to ~900 ml H2O. Add 1.14 ml glacial Acetic Acid and 2 ml

of 0.5 M EDTA solution (pH 8) and mix. Pour mixture into 1 L graduate cylinder and add H2O to a total volume of 1 L.

3. 6X Sample Loading Buffer: Combine 1 ml of sterile H2O and 1 ml of Glycerol. Add enough bromophenol blue powder to make the solution a deep blue color (about 0.05 mg).Note: Bromophenol blue migrates at a rate equivalent to 200–400bp DNA.

4. Ethidium Bromide (10 mg/ml)5. DNA ladder standard6. DNA fragments of different sizes7. Electrophoresis chamber8. Power supply9. Gel casting tray and combs


10. UV illuminator11. Gloves and goggles

Experimental procedure

Preparing the agarose gel

1. Measure 1 g Agarose powder and add it to a 500 ml flask.2. Add 100 ml TAE Buffer to the flask.3. Melt the agarose in a microwave or hot water bath until the solution becomes clear. (if

using a microwave, heat the solution for several short intervals - only until the solution starts to boil).

4. Let the solution cool to about 50-55°C, swirling the flask occasionally so it cools evenly.5. Place the combs in the gel casting tray.6. Pour the melted agarose solution into the casting tray and let cool until it is solid (it should

appear milky white).7. Carefully pull out the combs.8. Place the gel in the electrophoresis chamber.9. Add enough TAE Buffer so that there is about 2-3 mm of buffer over the gel.

Loading the gel

1. Add 10 l of each sample (DNA fragments) to 2 l of 6X Sample Loading Buffer (make sure the order which each sample is loaded has been recorded).

2. Carefully pipette each sample/Sample Loading Buffer mixture into separate wells in the gel.3. Pipette 5 l of the DNA ladder standard into another well of the gel.

Running the gel

1. Place the lid on the gel box, connecting the electrodes.2. Connect the electrode wires to the power supply, making sure the positive (red) and

negative (black) are correctly connected.3. Turn on the power supply.4. Check to make sure the current is running through the buffer by looking for bubbles

forming on each electrode.5. Check to make sure that the current is running in the correct direction by observing the

movement of the blue loading dye – this will take a couple of minutes (it will run in the same direction as the DNA).

6. Let the power run until the blue dye approaches the end of the gel.7. Turn off the power and disconnect the wires from the power supply.8. Remove the lid of the electrophoresis chamber.9. Carefully remove the gel


Staining the Gel

1. Wear gloves when working with Ethidium Bromide. 2. Place the gel in the staining tray containing 1.0 g/mL Ethidium Bromide solution. Allow

the gel to stain for 20-30 minutes.3. Remove the gel to a tray of water and allow to destain for 5 minutes.4. Place the gel on UV light source and observe the DNA bands.

Note: Never look/observe Ethidium Bromide fluorescence with unprotected eyes; short wave UV is damaging.

5. Take a picture.

Analysis

Label the identity of each sample by comparing the size of the bands on the DNA ladder standard.


bioanalytical chemistry (chem311)- lab manual

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