prepared bysite.iugaza.edu.ps/shashi/files/2011/09/practical...exercise 1 mitotic cell division...

Prepared by

Mazen M. El Zaharna

Prof. Fadel A. Sharief

First Edition 2008

A shy person does learn, nor should a

short-tempered person teach

Contents

Laboratory Safety 1

Exercise 1

Mitotic Cell Division 3

Exercise 2

Agarose Gel Electrophoresis 7

Exercise 3

Human Metaphase Chromosomes 13

Exercise 4

Human Chromosome Identification by G-Banding (karyotyping) 17

Exercise 5

Extraction of Human DNA 24

Exercise 6

Polymerase Chain Reaction (PCR) 28

Exercixe 7

Restriction Enzyme Digestion & Southern Blotting of DNA 32

Exercixe 8

Plasmid DNA Isolation 38

Exercixe 9

Transformation of Escherichia coli 41

Exercixe 10

SDS-PAGE 45

Practical Genetics 1st ed. 2008 1

Laboratory Safety Safety in the Laboratory Should always be in your mind. Throughout this manual Safety recommendations are given, below are some general consideration that anyone in a laboratory should know.

• General laboratory safety precaution. 1. Follow all instructions carefully. Use special care when you see the

word CAUTION in your laboratory instructions. Follow the safety instructions given by your instructor.

2. Determine the location of Fire Extinguishers, Chemical safety showers and Eye washers, Chemical Spill Kits, and alternative exit routes for lab evacuation.

3. Remember that smoking, eating, or drinking in the lab room is totally prohibited.

4. Lab coats should be always worn during the work in the lab. 5. Wear safety goggles when using dangerous chemicals, hot liquids, or

burners. 6. Any chemicals spilled on the hands or other parts of the skin should be

washed off immediately with a plenty of running water. 7. If you have an open skin wound, be sure that it is covered with a water

proof bandage. 8. Never work alone in the laboratory. 9. Keep your work area clean & dry. 10. Turn of all electrical equipment, water, and gas when it is not in use,

especially at the end of the laboratory period. 11. Report all chemicals spills or fluids to your instructor immediately for

proper clean up.

• Special precautions for working with heat or fire: 1. Never leave a lighted Bunsen burner unattended. When an object is

removed from the heat & left to cool, it should be placed where it is shielded from contact.

2. Inflammable liquid bottles should not be left open, not dispensed near a naked flame, hot electric element or electric motor.

3. Use test tube holders to handle hot laboratory equipments. 4. When you are heating something in a container such as a test tube,

always point the open end of the container away from yourself & others.

• Special precautions for working with chemicals 1. Never taste or touch substances in the laboratory without specific

instructions. 2. Never smell substances in the laboratory without specific instructions. 3. Use materials only from containers that are properly labeled. 4. Wash your hand after working with chemicals. 5. Do not add water to acid. Instead, dilute the acid by adding it to water.


• Special precautions for working with electrical equipment. 1. Make sure the area under & around the electrical equipment is dry. 2. Never touch electrical equipment with wet hands. 3. Make sure the area surrounding the electrical equipment is free of

flammable materials. 4. Turn off all power switches before plugging an appliance into an outlet.

• Special Precaution for working with Glasswares and other laboratory equipments.

1. Become familiar with the names and appearance of all the laboratory equipments you will use.

2. Never use broken or chipped glassware. 3. Make sure that all glasswares are clean before you using it. 4. Do not pick up broken glass with your bare hands. Use a pan and a

brush. 5. If a Mercury thermometer breaks, do not touch the mercury. Notify

your instructor immediately. 6. Use care handling all sharp equipments, such as scalpels and

dissecting needles.

• Special precautions for working with live or preserved specimens. 1. If live animals are used treat them gently. Follow instructions for their

proper care. 2. Always wash your hands after working with live or preserved

organisms. 3. Do not open Petri dishes containing live cultures unless you are

directed to do so. 4. Detergents (detol 5 – 10%) should be used to sterilize and clean

benches, glassware and equipment. 1. Safety cabinet should be used while working with microbes. 2. Disposable items should be collected and autoclaved.

Practical Genetics 1st ed. 2008

Exercise 1: Mitotic Cell Division

Objectives:

• Learn a staining procedure to identify the stages of mitosis in onion root tip.

• To differentiate between the different stages of mitosis.

• Calculate the mitotic index.

Introduction:

The growth and development of every organism depends on the precise replication of the genetic material during each cell division. Cell division is the process by which cells reproduce themselves, involves both division of the cell’s nucleus (karyokinesis) and division of the cytoplasm (cytokinesis). There are two types of nuclear division: mitosis and meiosis. New body (somatic) cells are formed by mitosis. Each cell division produces two new daughter cells with the same number and kind of chromosomes as the parent cell. The formation of male and female gametes in animal cells or spores in plant cells is by meiosis. Gametes and spores will have half the chromosome number of the parent cells.

Stages of mitosis

Interphase Interphase, which begins when cell division ends and continues until the beginning of the next round of division, is organized into three phases. G1, S and G2 (Figure 1.1). The chromatin in this phase is undifferentiated in the heavily-stained nucleus. Before the cell enters the mitosis phase, it first undergoes a synthesis or S phase where each chromosome is duplicated and consists of two sister chromatids joined together by a centromere. Centromeres are crucial to segregation of the daughter chromatids during mitosis. Now, the nucleus and cell increase in size, and chromosomes are fully extended. The cell is preparing for the beginning of mitosis. Mitosis Mitosis is the next phase of the cell cycle. It is essentially the same whether considering a simple plant or a highly evolved organism, such as a human being. The major function of mitosis is to accurately and precisely replicate genetic information, or chromosomes, so each daughter cell contains the same information. The process of mitosis is an ongoing event that can be segmented into several identifiable stages. In order, these stages are: prophase, metaphase, anaphase, and telophase. Cytokinesis (Figure 1.2), the actual process of cell division, occurs during telophase. In plants such as

Figure 1.1. Stages of the cell cycle.

1

Exercise 1 Mitotic Cell Division


the onion, this is seen as the formation of the cell plate between the two daughter cells. Prophase In prophase, dramatic changes begin to occur within the nucleus of the cell. Chromosomes become thicker, shorter, and easily visible when stained under the light microscope. Two “sister chromatids” join near their middle at a structure called the centromere. The nucleolus and the nuclear membrane disappear. The mitotic apparatus the spindle, begins to organize within the cell. Microtubules are slender rods of protein responsible for pulling replicated chromosomes towards each half of the cell. Metaphase During this period, chromosomes become aligned at midpoint or equator between poles of the cell and are at their thickest and shortest structure. They are easily identified as two longitudinally double sister chromatids. Chromatids are connected (at their centromeres) to the spindle apparatus, which has formed between the two centrioles located at the poles of the cell. In many plants, the centrioles are absent. The spindle is still present, however, and the plant chromosomes are similarly attached to the spindle microtubular fibers. Anaphase In this short phase, sister chromatids begin to separate and migrate to the poles. Once the two chromatids separate, each is called a chromosome. For humans, with a diploid number of 46 chromosomes, there will be 46 chromosomes moving toward each pole. Onions have 16 diploid chromosomes and, therefore 16 chromosomes move to each pole. During anaphase there is a quantitative, equal segregation of the diploid number of chromosomes into two developing nuclei at the poles of the anaphase cell. Telophase and Cytokinesis The final mitotic phase of the cell cycle is recognized by the formation of two new nuclei encompassing the daughter chromosome at the cell poles. The mitotic apparatus disappears and chromosomes begin to lengthen as they unwind. Cytokinesis, formation of a new cell membrane, occurs midway between the daughter nuclei. In plants, such as the onion root tip cells, this is seen as the formation of a cell plate, dividing the original cell into two (presumably equivalent) daughter cells. Cells now enter the G1, stage of interphase in the cell cycle and the process begins anew.

Prophase

Metaphase

Early anaphase

Late anaphase

Telophase

Figure 1.2. The phases of mitotic division.



Figure 1.3. The stages of mitosis in onion root tip cells.

In a growing plant root, the cells at the tip of the root are constantly dividing to allow the root to grow. Because each cell divides independently of the others, a root tip contains cells at different stages of the cell cycle. This makes a root tip an excellent tissue to study the stages of cell division. In this exercise, the student will prepare slides containing stained onion root tip squash sections, which will allow him to identify different stages of mitosis.

Procedure

Materials

� Slides & cover slips � Microscope � Fresh onion root tips � Fixative ( methanol-acetic acid 3:1 v/v) � Forceps � 1 M HCl � Razor blade � Stain � Paper towel, or absorbent paper

Method

Root tip preparation:

1. Obtain two small cups, label one of them with " HCl " and pour enough 1 M HCl into it to cover the bottom, likewise, label the other " fixative " and pour enough fixative fluid in it to cover the bottom.

Caution

Fixative (3:1 methanol to acetic acid) is a dangerous and volatile solution. Methanol is a flammable poison capable of causing skin irritation, blindness, CNS depression and death. Acetic acid is a flammable liquid capable of causing severe eye, skin, and respiratory chemical burns. Always wear personal protective equipm-ent, wash with plenty of water and move to fresh air if exposed.



2. Use forceps to transfer an onion root tip into the cup of HCl. After 4 minutes, transfer the root tip in the fixative and leave it for 4 minutes. Then place the root tip on a slide.

3. With a razor blade, or other sharp instrument cut off one to two mm of the root tip and discard all except the tip that you want to prepare.

4. Cover the root tip with a few drops of stain for 2 minutes, then blot away the stain. Be careful not to touch the root tip!

5. Cover the root tip with one to two drops of water put a cover slip over the root, put a paper towel or other absorbent paper and with your thumb firmly press on the cover slip.

Note: Do not twist the cover slip! This pressure will spread the cells into a single layer.

6. Observe your preparation under the low power (X10) of a microscope (Figure 1.4).

7. Search the slide to find cells in various stages of cell division, once you have located cells in division, change to high power (X40) & try to observe several stages of division.

8. Record the number of cells in each stage. Count at least three full fields of view. You should have counted over 200 cells.

9. Record your data in the table below. 1. Calculate the percentage of cells in each

phase and record in the table below.

Figure 1.4. Longitudinal section of onion root tip.

Number of

cells Percentage of total

cells counted Interphase Prophase

Metaphase Anaphase Telophase

Total Calculate the Mitotic Index

Number of cells in mitotic phase Mitotic Index =

Total number of cells counted


Exercise 2: Agarose Gel Electrophoresis

Objectives

• To understand the principle of Gel electrophoresis.

• To become familiar with the part of the electrophoresis setup.

Introduction

Electrophoresis is a laboratory technique for separating molecules based on their charge. Molecules with a negative charge (anions) will be attracted to the positively charged node (anode). Molecules with a positive charge (cations) will be attracted to the negatively charged node (cathode). Agarose gel electrophoresis is a widely used procedure in various areas of biotechnology. This simple, but precise, analytical procedure is used in research, biomedical and forensic laboratories. Of the various types of electrophoresis, agarose gel electrophoresis is one of the most common and widely used methods. It is a powerful separation method frequently used to analyze DNA fragments, and it is a convenient analytical method for determining the size of DNA molecules in the range of 500 to 30,000 base pairs. It can also be used to separate other charged biomolecules such as dyes, RNA and proteins. The separation medium is a gel made from agarose, which is a polysaccharide derivative of agar. Originating from seaweed, agarose is highly purified to remove impurities and charge. It is derived from the same seaweed as bacterial agar used in microbiology, as well as a food product called agar-agar. How Separation Occurs

The degree of separation and rate of molecular migration of molecules in a mixture depends upon two main factors, the electrical charge and the size of molecules (Figure 2.1).

Figure 2.1. Mixture of charged molecules can be separated into positive and negative ions due to the electric field, then they are separated according to their size due to gel.

Charge

Separation

Size

Separation

Analyze

Identify

Purify Mixture of

Charged Molecules

Positive Molecules

Negative Molecules

2

Exercise 2 Agarose Gel Electrophoresis


1- The electrical charge

In electrophoresis, the electric charge often is passed through what is known as a support medium. In general, the medium is mixed with a chemical mixture called a buffer. The buffer carries the electric charge that is applied to the system. The medium/buffer matrix is placed in a tray. Samples of molecules to be separated are loaded into wells or slots that have been formed at one end of the matrix. As electrical current is applied to the tray, the matrix takes on this charge and develops positively and negatively charged ends. As a result, molecules that are negatively charged such as deoxyribonucleic acid (DNA) and ribonucleic acid (RNA) are pulled toward the positive end of the gel. Proteins have net charges determined by charged groups of the amino acids from which they are constructed. Proteins can also be amphoteric compounds (a compound that can take on a negative or positive charge depending on the surrounding conditions). A protein in one solution might carry a positive charge in a particular medium and thus migrate toward the negative end of the matrix. In another solution the same protein might carry a negative charge and migrate toward the positive end of the matrix. For each protein there is a pH in which the protein molecule has no net charge (the isoelectric point). 2- Size of Molecules

The agarose gel consists of microscopic pores that act as a molecular sieve which separates molecules based upon size and shape. The charge to mass ratio is the same for different sized DNA molecules. Therefore, the absolute amount of charge on the molecule is not a critical factor in the separation process. The separation occurs because smaller molecules pass through the pores of the gel more easily than larger ones (Figure 2.2), i.e., the gel is sensitive to the physical size of the molecule. If the size of two fragments are similar or identical, they will migrate together in the gel.

Figure 2.2. Crosssectional area of agarose gel representing the pores present, small molecules move faster than large molecules. Photo at the left represents a scanning electron micrograph of agarose gel.

Molecules can have the same molecular weight and charge but different shapes, as in the case of plasmid DNAs. Molecules having a more compact shape (a sphere is more compact than a rod) can move more easily through the pores. The migration rate of linear fragments of DNA is inversely

Porous

Material Molecules

Entering Porous

Material

Smallest Move

Fastest



proportional to the log10

of their size in base pairs. This means that the

smaller the linear fragment, the faster it migrates through the gel. In addition, different molecules can interact with agarose to varying degrees. Molecules that bind more strongly to the agarose will migrate more slowly. The mobility of molecules during electrophoresis is also influenced by gel concentration. Higher percentage gels, as well as thicker gels, are sturdier and easier to handle. However, the mobility of molecules will take longer because of the tighter matrix of the gel (Figure 2.3).

Figure 2.3. "A" is a 1% agarose gel, the pores are bigger than "B" gel which is a 2% agarose gel.

In this exercise we will electrophorese various dyes which will approximate the physical properties of the negatively charged biomolecules ( eg. DNA, RNA and many proteins).

Procedure Materials

• Horizontal gel electrophoresis apparatus

• Agarose

• Electrophoresis buffer

• Direct Current (D.C.) power supply

• Automatic micropipettes with tips

• Balance

• Microwave or hot plate

• 250 ml flasks or beakers

• Distilled or deionized water

Method PREPARING THE GEL BED 1. Close off the open ends of a clean and dry gel bed

(casting tray) by using rubber dams. Place a rubber dam on each end of the bed. Make sure the rubber dam fits firmly in contact with the sides and bottom of the bed.

2. Place a well-former template (comb) in the first set of notches at the end of the bed. Make sure the comb sits firmly and evenly across the bed.

Electrophoresis Buffers

Depending on the size of the DNA electrophoresed and the application, diff-erent buffers can be used for agarose electrophoresis. TAE buffer (Tris Acetate EDTA) is the most common used agarose gel electrophoresis buffer. TAE has the lowest buffering capacity of the buffers, however TAE offers the best resolution for larger DNA. However, TAE requires a lower voltage and more time. However TBE buffer (Tris/Borate/EDTA) is often used for smaller DNA fragments (ie less than 500bp).

A B

1



CASTING AGAROSE GELS 3. Weigh the specified amount of agarose and transfer it to

a 250 ml flask. 4. Add the specified amount of buffer, swirl the mixture to

disperse clumps of agarose powder. 5. Heat the mixture to dissolve the agarose powder. The

final solution should appear clear (like water) without any undissolved particles.

� Microwave method:

Heat the mixture on High for 1 minute. Swirl the mixture and heat on High in bursts of 25 seconds until all the agarose is completely dissolved.

6. Cool the agarose solution to 55°C with careful swirling to promote even dissipation of heat.

7. Pour the cooled agarose solution into the bed. Make sure the bed is on a level surface. Allow the gel to completely solidify. It will become firm and cool after approximately 20 minutes.

Preparing the gel for electrophoresis 8. After the gel is completely solidified, carefully and

slowly remove the rubber dams from the gel bed. Be especially careful not to damage or tear the gel wells when removing the rubber dams.

9. Remove the comb by slowly pulling straight up. Do this carefully and evenly to prevent tearing the sample wells.

10. Place the gel (on its bed) into the electrophoresis chamber, properly oriented, centered and level on the platform.

11. Fill the electrophoresis apparatus chamber with the required volume of diluted buffer.

Loading the samples 12. Load each of the dye samples (A – E) into the wells in

consecutive order. The amount of sample that should be loaded is 10 µl.

Running the gel 13. After the samples are loaded, carefully put the cover

down onto the electrode terminals. Make sure that the negative and positive color-coded indicators on the cover and apparatus chamber are properly oriented.

14. Insert the plug of the black wire into the black input of the power source (negative input). Insert the plug of the red wire into the red input of the power source (positive input).

15. Set the power source at the required voltage and conduct electrophoresis for the length of time

1 2

3

4

5

6

7

9



determined by your instructor. General guidelines are presented in Table 1.

16. Check to see that current is flowing properly - you should see bubbles forming on the two platinum electrodes.

17. After approximately 10 minutes, you will begin to see separation of the colored dyes. After the electrophoresis is completed, turn off the power, unplug the power source, disconnect the leads and remove the cover.

18. Document the gel results. A variety of documentation methods can be used, including drawing a picture of the gel, taking a photograph, or scanning an image of the gel on a flatbed scanner.

Calculation and Analysis Measure the distance that each of the color bands migrated from the origin in the wells and record results in the table below. Rate of migration of a molecule is inversely proportional to the log of its molecular weight

Distance α 1 / log

10-MW

Construct a standard curve to relate the distance of migration of each dye to its assumed molecular weight. From this standard curve estimate the molecular weight of the unknown dye.

Dye Molecular Weight/ Dalton Distance migrated/ mm Cresol red 382.42 Bromophenol blue 669.96 Eosin 691.85 Orange G Trypan blue 960.81

12

13



Distance/ mm

Lo

g-

Mo

lecu

lar

we

ight


Exercise 3: Human Metaphase Chromosomes Objectives

� Preparing, staining and observing human metaphase chromosomes.

Introduction Chromosome Morphology Chromosomes are not visible under the light microscope in non-dividing (interphase) cells. As the cell begins to divide, the threads of chromatin (DNA-protein complex) in the nucleus begin to condense into multiple levels of coiled structures recognizable as chromosomes. There are two modes of cell division: mitosis and meiosis. Because mitotic cells are easy to obtain, morphological studies are generally based on mitotic metaphase chromosomes. Chromosome Structure At metaphase the chromosomes are at their most condensed state, with spindle fibers attaching to the area of the centromere called the kinetochore (Figure 3.1). Anaphase begins with the division of the centromere and the separation of chromatids. Once separated, each chromatid is known as a chromosome. The best mitotic stage for chromoso-me analysis is prometaphase or metaphase. A typic-al metaphase chromosome consists of two arms separated by a primary constriction or centromere. Each of the two sister-chromatids contains a highly coiled double helix of DNA. Often the sister chrom-atids are so close to each other that the whole chromosome appears as a single rod-like structure. A chromosome may be characterized by its total length and the position of its centromere (Figure 3.2). Chromosome Number The diploid chromosome number is the number of chromosomes in the somatic cell and is designated by the symbol 2N. The gametes, which have one-half the diploid number, have the haploid number N. In humans the diploid number is 46, with 23 inhe-rited from each parent through the sperm or egg. Same (homologous) chromosomes form a pair with one member from each parent. Thus, there are 23 pairs of chromosomes in human cells. Of these, 22 pairs are not directly involved in sex determination, and are known as autosomes. The remaining chromosome pair consists of the sex chromosomes, and is directly involved in sex determination. In females the two sex chromosomes are identical (XX),

Figure 3.1. Chromosomes are replicated during S phase, before mitosis begins. Two genetically identical chromatids of a replicated chromosome join at the centromere at the kinetochore area (a). In the photograph (b), a human chromosome is in the midst of forming two chromatids.

3

Exercise 3 Human Metaphase Chromosome


whereas in males the two sex chromosomes are not identical (XY).

Cytogenetics, the study of chromosomes, originated more than a century ago. However, not until the last 30 years or so have human chromosome studies become a major field in the biomedical sciences. Chromosome banding methods, for example, are today’s vital tools in clinical genetics and evolutionary studies. Furthermore, human cytogenetics coupled with molecular techniques has revolutionized the field of molecular genetics, including gene mapping and recombinant DNA technology.

Specimen In order to successfully culture blood samples for cytogenetic analysis they have to be received fresh and unclotted. Heparinized whole blood (green top vacutainer tube); sodium heparin is the recommended anticoagulant. Any other choice of tube is unsuitable since the sample will either arrive clotted or tube will contain EDTA which is toxic to cells and will therefore adversely affect the culture. Sterile Technique Aseptic or sterile technique is the performance of tissue culture procedures without introducing contaminating micro-organisms from the environment. In doing tissue culture work, 70% of the problems are due to a lack of good sterile technique. Microorganisms causing the contamination problems exist everywhere, on the surface of all objects and in the air. Tissue culture media used are often supplemented with antibiotics. Antibiotics do not eliminate problems of gross contamination which result from poor sterile technique or antibiotic-resistant mutants. Autoclaving renders pipettes, glassware, and solutions sterile. Overview of procedure for chromosome preparation A variety of tissue types can be used to obtain chromosome preparations. Some examples include peripheral blood, bone marrow, amniotic fluid, skin fibroblasts and products of conception. In the case of blood cell culture only cells that are actively dividing can be used for cytogenetic studies. Normally only white blood cells are used for cytogenetic analysis (i.e. neutrophils, eosinophils, basophils, monocytes and lymphoctes). These cells contain a nucleus and are capable of undergoing cell division. Although specific techniques differ according to the type of tissue used, the basic method for obtaining chromosome preparations is as follows:

1- Cell culture: although cell culture methods vary significantly with the tissue of origin (amniotic fluid, chorionic villi, fetal tissues, peripheral blood, bone marrow, solid tumors, cell lines of various origins), the final goal is to achieve cell growth and division, ultimately leading to a good mitotic index.

Figure 3.2. Chromosome morphology and terminology.



2- Harvesting: mitotic spindle formation is blocked, usually by adding colcemide to the culture, and the cell division is stopped at the metaphase level. Cells are subjected to hypotonic treatment, which increases their volume, and disrupts the cell membrane of the red blood cells allowing their removal. A fixative solution is added to the cell suspension to preserve the cells in their "swollen" state and to remove the water, thus "hardening" the biologic material. The common fixative (3:1 methanol:acetic acid) removes lipids and alters/denatures proteins thus making the cell membrane remnant very fragile, which is important for subsequent chromosome spreading.

3- Slide preparation: drops of cell suspension are placed on a slide, and allowed to dry in a controlled fashion, leading to chromosome spreading.

4- Staining: stain chromosome preparations to detect possible numerical and structural changes.

Procedure

Materials � Pre-cleaned microscope slides � 5 ml peripheral blood in sodium heparin (500 U/ml) � 0.04 mg/ml colchicine � Lymphocyte culture medium: RPMI 1640, 100000 U/ml penicillin, 100

mg/ml streptomycin, 20% fetal bovine serum (FCS), Phytohem-agglutinin 20µg/ml

� Hypotonic solution: 0.075 M KCl � Fixative: 3:1 methanol/ acetic acid at 4°C

Method

Cell culture 1. Label a 15 ml sterile culture tube, pipette 10 ml RPMI 1640 medium with L-

Glutamine. Add the following supplements to the tube: 10 µl Penicillin-streptomycin, 0.3 ml Phytohemagglutinin and 2 ml of Fetal bovine serum.

2. Add 1 ml of heparinized blood into the medium tube. 3. Mix the contents of the tube with gentle inversion, then incubate the tube in

5% CO2 incubator at 37°C for 72 hours in a slant position.

Harvesting

a- Stopping cell division at metaphase

4. After 72 hr culture, add 25 µl of pre-warmed colchicine (37C) and mix well gently. Then incubate the tube for 30 min at 37°C.

b- Hypotonic treatment of cells 5. Centrifuge tubes at 2000 rpm for 10 min.

Caution Fixative (3:1 methanol to acetic acid) is a dangerous and volatile solution. Methanol is a flammable poison capable of causing skin irritation, blindness, CNS depression and death. Acetic acid is a flammable liquid capable of causing severe eye, skin, and respiratory chemical burns. Always wear personal protective equipment, wash with plenty of water and move to fresh air if exposed.



6. Discard supernatant without disturbing the cells on the bottom leaving about 0.5 ml of medium above the cell pellet.

7. Resuspend the cells in the remaining medium and carefully add 1 ml of prewarmed (37°C) 0.075 M KCl, drop-by-drop, while agitating gently. Add an additional 9 ml of KCl, for a total of 10 ml; mix well.

8. Incubate for 15 min at 37°C in the incubator. c- Fixation of Cells 9. Centrifuge tube at 1500 rpm for 10 minutes. 10. Remove supernatant leaving about 0.5 ml of fluid above the cells. 11. Add 5 ml of freshly prepared fixative, the first 2 ml should be added

dropwise while agitating gently, recap the tube and invert to mix. 12. Incubate the tube in the refrigerator for 30 minutes. 13. Centrifuge the tube at 1500 rpm for 10 minutes. 14. Remove the supernatant and add 6 ml of fixative, mix well and then

centrifuge at 1500 rpm for 10 minutes. 15. Repeat step 14 one or two times as needed. 16. Remove supernatant leaving about 1 ml of fixative, resuspend cells. This

is the material which is going to be used for slide preparation. d- Slide Preparation and Staining The slides should be exceptionally clean. 17. Withdraw a few drops of the cell suspension with a Pasteur pipette. 18. From a height of approximately 20 cm drop 2 or 3 drops of fluid onto each.

slide. 19. Allow the slides to dry, the best way is to place the slide in an incubator at

37°C overnight 20. Stain the slides by immersion in freshly prepared Giemsa stain for 7-10

minutes. 21. Remove slides from stain and rinse in distilled water until all excess satin

is removed. e- Observing human metaphase chromosomes Observe the slides under the microscope, first locate the field under low power (X10) lens, then observe the slide under high power (X40) lens and determine the best metaphase spread. Now observe the slide under oil immersion lens, and start examination of different chromosomes.

Figure 3.3. "a" represents a metaphase spread under high power lens, "b" represents a metaphase spread under oil immersion lens.

Important Note Hypotonic solution should not remain in contact with the cells for more than 27 minutes. Excess exposure may cause

rupture of WBCs.

a b


Exercise 4: Human Chromosome Identification by G-Banding (karyotyping)

Objectives

• Preparation, Staining and Observing G-banded human chromosomes. • Develop an understanding of karyotyping and the association of

various chromosomal abnormalities to diseases. Introduction Chromosomes are composed of double-stranded DNA associated with specific proteins. The nuclei of normal human somatic cells each contain 23 pairs of chromosomes, one of each pair derived from either maternal or paternal origin. During metaphase, chromosomes become condensed and stain intensely with basic dyes. The size and staining patterns allow the differentiation of different chromosomes. The autosomes, or nonsex chromosomes, are numbered from 1 to 22 approximating decreasing size order. The 23rd pair is the sex chromosomes, X and Y; normal females contain two X chromosomes per nucleus, whereas normal males contain one X and one Y chromosome. Variations in the karyotype, can be used to diagnose certain disorders (e.g. Down syndrome), prenatal abnormalities or certain types of tumors. A karyotype is a display or photomicrograph of an individual’s somatic-cell metaphase chromosomes that are arranged in a standard sequence, usually based on number, size, and type (Figure 4.1). Chromosome abnormalities Chromosome abnormalities can be numerical, as in the presence of extra or missing chromosomes, or structural as in translocations, inversions, large scale deletions or duplications. Numerical abnormalities, also known as aneuploidy, often occur as a result of nondisjunction during meiosis in the formation of a gamete; trisomies, in which three copies of chromosome are present instead of the usual two, are common numerical abnormalities.

Some chromosomal abnormalities that lead to disease in humans include:

• Turner syndrome results from a single X chromosome (45, X or 45, X0). • Klinefelter syndrome, the most common male chromosomal disease,

otherwise known as 47, XXY is caused by an extra X chromosome. • Down syndrome, a common chromosomal disease, is caused by trisomy

of chromosome 21.

Some disorders arise from loss of just a piece of one chromosome, including:

Figure 4.1. Karyotype of a normal male.

4

Exercise 4 Human Chromosome Identification by G-Banding


• Cri du chat (cry of the cat), from a truncated short arm on chromosome 5. The name comes from the babies' distinctive cry, caused by abnormal formation of the larynx.

• 1p36 Deletion syndrome, from the loss of part of the short arm of chromosome 1.

• Angelman syndrome – 50% of cases have a segment of the long arm of chromosome 15 missing.

Chromosomal abnormalities can also occur in cancerous cells of an otherwise genetically normal individual; one well-documented example is the Philadelphia chromosome, a translocation mutation commonly associated with chronic myeloid leukemia and less often with acute lymphoblastic leukemia.

Identification of Chromosomes The pairs of chromosomes are differentiated according to the following characteristics (Figure 4.2):

� Size: This is the easiest way to tell two different chromosomes apart.

� Banding pattern: The size and location of Giemsa bands on chromosomes make each chromosome pair unique.

� Centromere position: Centromeres are regions in chromosomes that appear as a constriction.

Using these key features, scientists match up the 23 pairs. Performing a Karyotype The slides are scanned for metaphase spreads and usually 10 to 30 cells are analyzed under the microscope by a cytogeneticist. When a good spread (minimum number of overlapping chromosomes) is found, a photograph is taken or the analysis is done by a computer. The chromosomes are arranged in a standard presentation format of longest to shortest. Actually chromosome 21 is smaller than chromosome 22, however, since Trisomy 21 (Down Syndrome) had already been named, it was decided to leave the numbering system as it was. Centromere position The centromere is the location of spindle attachment and is an integral part of the chromosome. It is essential for the normal movement and segregation of chromosomes during cell division. Human metaphase chromosomes come in three basic shapes and can be categorized according to the length of the short and long arms and also the centromere location (Figure 4.3). Metacentric chromosomes have short and long arms of roughly equal length with the centromere in the middle. Submetacentric chromosomes have short and long arms of unequal length with the centromere more towards one end. Acrocentric chromosomes have a centromere very near to one end and have very small short arms. They frequently have secondary constrictions on the short arms that connect very small pieces of DNA, called stalks and satellites, to the centromere.

Figure 4.2. key features to identify chromosome similarities and diff-errences.



Figure 4.3. Classification of chromosomes according to the centromere position.

Banding patterns

Special chromosome-staining procedures have revealed specific sets of intricate bands (transverse stripes) in many different organisms. The positions and sizes of the bands are highly chromosome specific. One of the basic chromosomal banding patterns is that produced by Giemsa reagent, a DNA stain applied after mild proteolytic digestion of the chromosomes. This reagent produces patterns of light-staining (G-light) regions and dark-staining (G-dark) regions. The patterns are consistent within species. In the complete set of 23 human chromosomes, there are approximately 550 G-dark bands visible at metaphase of mitosis. These bands have provided a useful way of subdividing the various regions of chromosomes, and each band has been assigned a specific number.

The difference between dark- and light-staining regions was believed to be caused by differences in the relative proportions of bases: the G-light bands being relatively GC-rich, and the G-dark bands AT-rich. However, it is now thought that the differences are too small to account for banding patterns. The crucial factor appears to be chromatin packing density: the G-dark regions are packed more densely, with tighter coils, which results in a higher density of DNA to take up the stain.

In addition, various other correlations have been made. For example, deoxynucleotide-labeling studies showed that G-light bands are early replicating. Furthermore, if polysomal (polyribosomal) mRNA (representing genes being actively transcribed) is used to label chromosomes in situ, then most label binds to the G-light regions, suggesting that these regions contain most of the active genes. From such an analysis, it was presumed that the density of active genes is higher in the G-light bands.



Procedure Materials

� Hank’s Balanced Salt Solution (HBSS) � Giemsa stain � Phosphate Buffer, pH 6.8 � Normal saline (0.9%) � 0.25% Trypsin (working solution is prepared by mixing 5 ml of 0.25%

Trypsin with 45 ml normal saline) Method We are going to follow the same procedure indicated in exercise 3 till we prepare the slides, and they will be ready for staining. Method for obtaining chromosome preparations is as follows:

• Cell culture,

• Harvesting,

• And Slide preparation. Age slides

� Place fixed dry slides on slide rack in 95oC oven and bake for 20 minutes. � Take slides out of the oven and leave them to cool.

Trypsinize slides with metaphase chromosomes � Immerse aged slides in working Trypsin Banding Solution at room

temperature for 15-120 seconds with a forceps. � Follow the Trypsin treatment with a brief immersion and swirl in phosphate

buffer in a coplin jar to stop the action of the Trypsin. Stain slides in Giemsa stain � Immerse the Trypsinized slides in working Giemsa stain for 2 minutes. � Rinse slides thoroughly with distilled water. � Tap off excess water on a paper towel, wipe the back of the slide dry and

leave the slide to dry. Observe the slides under the microscope, first locate the field under low power (X10) lens, then observe the slide under high power (X40) lens and determine the best metaphase spread. Now observe the slide under oil immersion lens, and start identification of different chromosomes. Procedure notes � Time of trypsin treatment varies with each case and is dependent on the

age of the slides to be banded, the technique used to make the slides, and the temperature and concentration of the trypsin solution.

� Chromosomes are under-trypsinized if they stain homogenously dark, banding is present but the bands appear to blur into each other, or if banding is present but a darkly stained bar is noticeable between the chromatids.

� Chromosomes are over-trypsinized if they appear swollen, fuzzy, or have a cobwebbed appearance.



� Due to variation in the stages of condensation of chromosomes on the same slide, some spreads will be over-trypsinized and some will be under-trypsined. The goal is to maximize the number of analyzable metaphases for a given slide.

� It is important to mix the working Leishman’s stain fresh before using, and change the solution if it has been sitting for more than approximately 30 minutes.

� Slides that are under-Trypsinized may be destained in fixative (3:1 methanol to acetic acid), rinsed in water, re-Trypsinized for a few seconds, and then re-stained.

Cut and Paste Cut the chromosomes present on page number 23 and paste each chromosome near its homologous pair on page 22. Now, indicate if the karyotype is normal or abnormal, if abnormal; indicate the abnormality present



X or YX or YX or YX or Y


Exercise 5: Extraction of Human DNA Objectives

• Isolation of genomic DNA from human blood. • Analysis of isolated DNA using agarose gel electrophoresis and

spectrophotometery. Deoxyribonucleic Acid (DNA) A nucleic acid that carries the genetic information in the cell and is capable of self-replication and synthesis of RNA. DNA consists of two long chains of nucleotides twisted into a double helix and joined by hydrogen bonds between the complementary bases adenine and thymine or cytosine and guanine (Figure 5.1). The sequence of nucleotides determines individual hereditary characteristics. DNA plays an important role in two processes. During the process of replication, DNA provides information to copy itself, so genetic information can be passed on from generation to generation of cells. DNA also provides instructions for making proteins, which are vital to the maintenance and function of cells. DNA provides information for the order of amino acids required for making various proteins. A large number of proteins and enzymes including DNA polymerases are involved in the synthesis of DNA. The human genome consists of about 2.9 billion base pairs. Of this total, only about 5% code for protein. Intervening sequences and other noncoding sequences make up the remainder. Some of the noncoding sequences or other functionally unassigned sequences may possess undiscovered functions. In addition to genomic DNA, mitochondria, which are cellular organelles, contain their own DNA (mitochondrial DNA) which replicate independently from cell chromosomal DNA. When cells are chemically lysed (broken open), DNA from chromosomes is released and can be isolated and purified. DNA extraction is frequently the first step for molecular biology and biotechnology experiments. Extracted DNA is soluble in water and thus will appear as a clear solution. By contrast DNA is insoluble in salt solutions and alcohol, where it will form white fibers. Purification procedures for DNA usually include precipitation with alcohol in the presence of salt. A glass rod or a stirrer is used to spool DNA and to separate the DNA from the solution. The DNA will appear as a viscous, clotted mass. The amount of DNA spooled will vary and is a consequence of the intactness of the DNA sample.

Figure 5.1. The DNA double helix.

5

Exercise 5 Extraction of Human DNA


Almost any tissue or body fluid may be used as a source of DNA. The most common sources of human DNA are samples from hair, cheek cells, blood and saliva. Once extracted, DNA can be stored for long periods of time. Various methods of storage include precipitation and storage under alcohol at room temperature or under refrigeration. DNA can be recovered from small blood spots and tissues, or even a few cells. Such amounts of DNA can be obtained from individuals during medical procedures or evidence left behind in crime scenes such as cells that are recovered from the fingernails of a victim. In recent times, a few cells deposited by a person while licking and sealing an envelope has been sufficient to obtain DNA and match the DNA fingerprint to the person who left this evidence behind. There are three basic steps in a DNA extraction, the details of which may vary depending on the type of sample and any substance that may interfere with the extraction and subsequent analysis.

• Break open cells and remove membrane lipids.

• Remove cellular and histone proteins bound to the DNA, by adding a protease, by precipitation with sodium or ammonium acetate, or by using a phenol/chloroform extraction step.

• Precipitate DNA in cold ethanol or isopropanol, DNA is insoluble in alcohol and clings together, this step also removes salts.

Analysis of the isolated DNA The product of DNA extraction will be used in subsequent experiments. Poor quality DNA will not perform well in PCR. Quality and quantity of the extracted DNA can be analyzed by 2 methods:

• agarose gel electrophoresis, where a good quality DNA is represented by a sharp band near the wells of the gel, while smearing indicates DNA degradation (Figure 5.2),

• and spectrophotometery for quantitation and purity.

Figure 5.2. A Photograph representing the quality of 16 DNA samples extracted from whole blood samples, run on ethidium bromide stained 1% agarose gel. Lanes 1,3,5,7,9,11,13 & 15 represent good quality DNA, while lanes 2, 4, 6, 8, 10, 12, 14 & 16 represent bad quality (broken) DNA.



Procedure Blood samples should be collected in disodium EDTA tube. Samples can be

stored at -20oC or -70oC. Fresh samples are kept in freezer for a few hours to facilitate RBCs hemolysis. Allow samples to thaw before starting the extraction. Materials:

• Erythrocyte lysing buffer (0.155M NH4Cl, 10 mM KHCO3, 0.1 mM Na2EDTA, pH 7.4)

• SE buffer (75 mM NaCl, 25 mM Na2EDTA, pH 8.0) containing 100

µg/ml of Proteinase K & 1% sodium dodecyl sulphate (SDS)

• 6 M NaCl

• Chloroform

• Isopropanol

• 75% ethanol

• TE buffer (1 M Tris-HCl; 0.5 M EDTA; pH 8.0) 1. Lysis of red blood cells

• Pipette 3 mls of whole blood in a conical centrifuge tube

• Add 9 mls of 1X erythrocyte lysing buffer

• Leave 10 minutes at RT, mix occasionally

• Centrifuge at 4000 rpm for 5 min

• Discard supernatent

• White pellet is observed at bottom of tube

• Wash pellet 3 times by adding 3 mls of buffer, incubate 10 min at RT, & centrifuge

2- Lysis of leukocytes and leukocytes' nuclei

• Add 1.5 mls of SE buffer to the pellet

• Incubate at 37-55oC overnight in a water bath or incubator

• WBCs denatured, nuclei are lysed & DNA goes out in solution 3- Extraction of proteins

• After the incubation, add 1.5 ml of SE buffer together with 750 µl of 6 M NaCl and then add 3.75 ml chloroform.

• The tubes are mixed vigorously (on vortex) for 20 seconds with occasional mixing for at least 30 minutes

• Centrifuge for 10 minutes at 2000 rpm with minimal breaking force

• After centrifugation 2 phases are observed and care must be taken not to disturb the interphase

• The upper phase contains the DNA while proteins are in the lower phase

4- DNA precipitation and wash

• The upper phase containing the DNA is transferred to a clean and sterile conical centrifuge tube



• Add an equal volume of isopropanol

• DNA will be precipitated by gentle swirling & observed as a white thread like strand

• Using a sterile spatula or loop transfer the DNA strand into a sterile microcentrifuge tube containing 1 ml of 75% ethanol

• Wash DNA by inversion to remove any remaining salts

• Centrifuge at 11000 g for 4 minutes, then discard supernatant taking care not to discard the pellet

• Repeat the washing step, then centrifuge

• Remove supernatant, and dry the pellet by using a vacuum centrifuge

or by leaving the tubes opened and inverted in an oven at 50-65oC for 1 hour

5- Resuspension of DNA

• Dried pellet is resuspended in TE buffer and left overnight on a rotator

6- Check quantity and quality DNA Quality and quantity of the extracted DNA can be analyzed by 2 methods: 1- Agarose gel electrophoesis Prepare agarose gel of 0.6% to 1% concentration, mix DNA samples with loading buffer and then load on the gel. Electrophorese at 70–80 volts, for 45–90 minutes. After electrophoresis stain the gel with ethidium bromide and then view on UV transilluminator. A good quality DNA is represented by a sharp band near the wells of the gel, while smearing indicates DNA degradation. 2- Spectrophotometry Spectrophotometer is used to measure the quantity and purity of the extracted DNA. Quantity: Nucleic acids have a peak absorbance in the ultraviolet range at about 260 nm

1 A260 O.D. unit for dsDNA = 50 µg/ml DNA purity: The purity of the DNA is reflected in the OD260:OD 280 ratio and

must be between 1.7 and 2.00. Decreased 260:280 ratio means that too much protein or other contaminant is present.


Exercise 6: Polymerase Chain Reaction (PCR) Objectives

• To understand how the PCR technique works • To perform the PCR experiment • To analyze the PCR products

Introduction The Polymerase Chain Reaction (PCR) is a powerful and sensitive technique for DNA amplification in vitro. PCR amplifies specific DNA sequences exponentially by using multiple cycles of a three-step process. PCR can achieve more sensitive detection and higher levels of amplification of specific sequences in less time than previously used methods, e.g. DNA cloning. These features make the technique extremely useful, not only in basic research, but also in commercial uses, including genetic identity testing, forensics, industrial quality control and in vitro diagnostics. Basic PCR has become commonplace in many molecular biology labs where it is used to amplify DNA fragments and detect DNA or RNA sequences within a cell or environment. However, PCR has evolved far beyond simple amplification and detection, and many extensions of the original PCR method have been described. PCR is capable of amplifying sequences from minute amounts of target DNA, even the DNA from a single cell. Such exquisite sensitivity has afforded new methods of studying molecular pathogenesis and has found numerous applications in forensic science, and in diagnosis, in genetic linkage analysis using single-sperm typing. However, the extreme sensitivity of the method means that great care has to be taken to avoid contamination of the sample under investigation by external DNA, such as from minute amounts of cells from the operator. The cycling reactions There are three major steps in a PCR (Figure 6.1), which are repeated for 30 or 40 cycles. This is done on an automated cycler, which can heat and cool the tubes with the reaction mixture in a very short time.

1. Denaturation The initial step denatures the target DNA by heating it to 94°C or higher for 15 seconds to 2 minutes. In the denaturation process, the two intertwined strands of DNA separate from one another, producing the necessary single-stranded DNA template for replication by the thermostable DNA polymerase.

2. Annealing

In this step the temperature is reduced to approximately 50–60°C. At this temperature, the oligonucleotide primers can form stable associations

Nucleic Acid Cross-Contamination

It is important to minimize cross-contamination between samples and prevent carryover of RNA and DNA from one experiment to the next. Use separate work areas and pipettes for pre- and post-amplification steps. Use aerosol-resistant tips to reduce cross-contamination during pipetting. Wear gloves and change them often.

6

Exercise 6 Polymerase Chain Reaction (PCR)


(anneal) with the denatured target DNA and serve as primers for the DNA polymerase.

3. Extension

Finally, the synthesis of new DNA begins as the reaction temperature is raised to the optimum for the DNA polymerase. For most thermostable DNA polymerases, this temperature is in the range of 70–74°C. The extension step lasts approximately 1–2 minutes. The next cycle begins with a return to 94°C for denaturation.

Denaturation step: An initial denaturation step of 2–5 minutes at 94–95°C is required prior to start PCR cycling to fully denature the DNA.

Figure 6.1. Schematic diagram of the PCR process.

PCR Reaction Components Typical components of a PCR include:

� DNA: the template used to synthesize new DNA strands. Use of high quality, purified DNA templates greatly enhances the success of PCR



reactions. It is also critical that contamination from outside sources,

especially previous PCR reactions, be avoided. Approximately 104 copies of the target DNA are required to detect a product in 25–30 cycles of PCR. Typically, this means a 1–10 µg/ml of genomic templates.

� DNA polymerase: an enzyme that synthesizes new DNA strands. Taq DNA polymerase is isolated from Thermus aquaticus and catalyzes the primer-dependent incorporation of nucleotides into duplex DNA in the 5′→3′ direction in the presence of Mg2+. It is recommended to use 1–1.25 units of Taq DNA polymerase in a 50µl amplification reaction.

� Two PCR primers: short DNA molecules (oligonucleotides) that define the DNA sequence to be amplified. Oligonucleotide primers are generally 20–30 nucleotides in length, and ideally have a GC content of 40–60%, with GC residues spaced evenly within the primer. Calculated melting temperatures (Tm) for the two primers should be from 42–65°C, and the Tm for the two primers should be within 5°C of each other.

� Deoxynucleotide triphosphates (dNTPs): the building blocks for the newly synthesized DNA strands. The final concentration of dNTPs is typically 200 µM of each nucleotide.

� Reaction buffer: a chemical solution that provides the optimal environmental conditions. Most reaction buffers consist of a buffering agent, most often a Tris-based buffer, and salt, commonly KCl. The buffer regulates the pH of the reaction, which affects the DNA polymerase activity and fidelity.

� Magnesium: a necessary cofactor for DNA polymerase activity. A magnesium concentration of 1.5–2.0 mM is optimal for most PCR products generated with Taq DNA Polymerase. Optimization normally involves supplementing the magnesium concentration in 0.5 or 1.0 mM increments.

Procedure Materials

� Standard Taq Reaction Buffer (10X) � Deoxynucleotide Solution Mix (10 mM) � Upstream Primer (10 µM stock) � Downstream Primer (10 µM stock) � DNA Template � Taq DNA Polymerase � Nuclease free water

Methods In this experiment we are going to amplify part of the Human Growth Hormone gene. Specific primers for this gene will to be used. 1. Prepare the master mix

When setting up multiple reactions it is faster and more accurate to create a master mix of the components that are common to all reactions. In



general, this involves creating a stock solution of polymerase, nucleotides, reaction buffer, water, and primers. The master mix is then aliquotted and mixed with the DNA template.

PCR reaction mixture

Reagent Volume

(µl) Final

concentration Volume X

No. of tubes

Buffer (X10) 2.0 1X

MgCl2 (25 mM) 1.6 2.0 mM

dNTPs (100mM) 0.1 0.1 mM

Primer 1 (F) 0.2 1.0 µM

Primer 2 (R) 0.2 1.0 µM

Taq DNA polymerase 0.25 2.0 U

DNA template 2.0 100 ng

Water 13.65

Total Volume 20

2- Program the thermocycler

The thermocycler profile for amplifying human growth hormone gene is:

� Step 1: Denaturation for 3 min. at 95oC

� Melting for 60 sec. at 95oC

� Annealing for 60 sec. at 57oC

� Extension for 90 sec. at 72oC

� Step 3: Final elongation for 10 min. at 72oC

� Step 4: Hold at 4 oC 3- Run the samples on thermocycler

Place the samples in the thermocycler, close the lid and start the program.

4- Analysis of PCR products � Analyze products on 2% agarose gel containing

ethidium bromide (Ethidium Bromide, A fluorescent dye visualized when excited by UV light).

� Loading Dye Samples are prepared with loading dye and then loaded on the gel. It is used to prepare DNA markers and samples for loading on agarose gels. It contains bromophenol blue dye, for visual tracking of DNA migration during electrophoresis. The presence of glycerol in the solution ensures that the sample sinks at the bottom of the well.

Visualize the PCR product on UV transilluminator, There should be a 400 bp band for the positive samples.

Important Note Negative & positive control reactions should be run with patient samples. Negative control to check for contamination, contains all reagents except the DNA template. Positive control to check if PCR reaction has worked, contains all reagents and a known target-containing DNA template.

Caution Ethidium bromide is a powerful mutagen and is toxic. Avoid breathing the dust. Wear appropriate gloves when working with solutions that contain this dye.


Exercixe 7: Restriction Enzyme Digestion & Southern Blotting of DNA Objectives

� Digestion of DNA by a restriction enzyme. � Analysis of the digested DNA by electrophoresis. � Transfer the digested DNA to nitrocellulose membrane (Southern blotting)

Including the procedure of setting up the Southern blotting device. Introduction The discovery of restriction enzymes ushered in a new era of molecular genetics. These enzymes cut the DNA molecule in a highly specific and reproducible way. This, in turn, has lead to the development of molecular cloning and the mapping of genetic structures. Restriction enzymes are endonucleases which catalyze the cleavage of the

phosphodiester bonds within both strands of DNA. They require Mg+2 for activity and generate a 5 prime (5') phosphate and a 3 prime (3') hydroxyl group at the point of cleavage. The distinguishing feature of restriction enzymes is that they only cut at very specific sequences of bases, Figure 7.1. Restriction enzymes are obtained from many different species of bacteria. To date, over 3,000 restriction enzymes have been discovered and catalogued. Restriction enzyme is part of the cell’s restriction-modification system in bact-eria. The phenomenon of restriction mod-ification in bacteria is a small scale immune system for protection from infection by foreign DNA. Bacteria can protect themselves only after foreign DNA has entered their cytoplasm. For this protection, many bacteria specifically mark their own DNA by methylating bases on particular sequences with modifying enzymes. DNA that is recognized as foreign by its lack of methyl groups on these same sequences is cleaved by the restriction enzymes and then degraded by exonucleases to nucleotides. Restriction enzymes are named according to the organism from which they are isolated. This is done by using the first letter of the genus followed by the first two letters of the species. Only certain strains or sub-strains of a particular species may produce restriction enzymes. The type of strain or substrain sometimes follows the species designation in the name. Finally, a Roman numeral is always

Figure 7.1. The enzyme EcoRI cutting DNA at its recognition sequence.

7

Exercixe 7 Restriction Enzyme Digestion & Southern Blotting of DNA


used to designate one out of possibly several different restriction enzymes produced by the same organism or by different substrains of the same strain. A restriction enzyme requires a specific double stranded recognition sequence of nucleotides to cut DNA. Recognition sites are usually 4 to 8 base pairs in length. Cleavage occurs within or near the site. The cleavage positions are indicated by arrows. Recognition sites are frequently symmetrical, i.e., both DNA strands in the site have the same base sequence when read 5' to 3'. Such sequences are called palindromes. Consider the recognition site and cleavage pattern of Eco RI as an example.

As shown above, Eco RI causes cleavage of DNA. The ends of the DNA fragments are called “sticky” or “cohesive” ends because the single-stranded regions of the ends are complementary. Other restriction enzymes, such as Hae III, introduce cuts that are opposite each other. This type of cleavage generates “blunt” ends, as shown below.

In general, the longer the DNA molecule, the greater the probability that a given recognition site will occur. Therefore, human chromosomal DNA, which contains three billion base pairs, has many more recognition sites than a plasmid DNA containing only several thousand base pairs. Restriction enzymes are traditionally classified into three types on the basis of subunit composition, cleavage position, sequence-specificity and cofactor-requirements. However, amino acid sequencing has uncovered extraordinary variety among restriction enzymes and revealed that at the molecular level there are many more than three different kinds. Type II enzymes cut DNA at defined positions close to or within their recognition sequences. They produce discrete restriction fragments and distinct gel banding patterns, and they are the only class used in the laboratory for DNA analysis and gene cloning. Unit Determination Assay It is important to know the unit determination assay of the enzyme, one unit of restriction endonuclease is defined as the amount of enzyme required to digest one microgram of the appropriate substrate DNA completely in 60 minutes under the conditions specified for that enzyme.



Digestion by Restriction Enzyme Materials

• DNA sample

• HinfI Restriction Enzyme

• Restriction enzyme buffer mix

• Nuclease-free water

Methods Measure the DNA concentration Use the Nano-drop spectrophotometer to measure the concentration of DNA, this is used to determine the amount of HinfI restriction enzyme to be used. Digestion of DNA Mix the following components in a clean microtube

Component Volume

nuclease-free water 16 µl

10X Buffer R 2 µl

DNA (0.5-1 µg/µl) 1 µl

HinfI (10 u/µl) 0.5-2 µl *

* The amount of the enzyme to be used depends on the concentration of DNA.

� Mix gently and spin down for a few seconds. � Incubate at 37°C for 16 hours.

Analysis of DNA digestion

� Analyze products on 2% agarose gel containing ethidium bromide. � Samples are prepared with loading dye and then loaded on the gel. � Visualize the PCR product on UV transilluminator.

Undigested DNA is represented by a sharp band near the wells of the gel, while smearing indicates digested DNA sample.



Southern Blotting A technique used in molecular biology to check for the presence of a particular DNA sequence in a DNA sample. The technique was developed by E.M. Southern in 1975. Southern Blotting could be used to locate a particular gene within an entire genome. The amount of DNA needed for this technique is dependent on the size and specific activity of the probe.

The total cellular DNA of an organism (genome) or the cellular content of RNA are complex mixtures of different nucleic acid sequences. Restriction digest of a complex genome can generate millions of specific restriction fragments and there can be several fragments of exactly the same size which will not be separated from each other by electrophoresis. Southern Blotting techniques have been devised to identify specific nucleic acids in these complex mixtures The Southern Blot takes advantage of the fact that DNA fragments will stick to a nylon or nitrocellulose membrane. DNA molecules are first elctrophoresed and then transferred from an agarose gel onto a membrane The membrane is laid on top of the agarose gel and absorbent material (e.g. paper towels or a sponge) is placed on top. With time, the DNA fragments will travel from the gel to the membrane by capillary action as surrounding liquid is drawn up to the absorbent material on top. The membrane is now a mirror image of the agarose gel.

Procedure Materials

• Whatman 3 mm Blotting Paper

• Nitrocellulose or nylon membrane filter

• Paper towels

• A weight

• 20x SSC (3M NaCl, 0.3M NaCitrate pH7.4) • 2x SSC

• 6x SSC

• Labeled probe Methods

Digest the DNA DNA is digested as indicated before with Hinf1 restriction enzyme. Electrophoresis Load digested samples onto agarose gel (typically 0.8 to 1.0% agarose). Run gel at maximum rate of 5 volts per cm. Denature the DNA (usually while it is still on the gel).



Soak it in about 0.5M NaOH, which would separate double-stranded DNA into single-stranded DNA. Only ssDNA can transfer. Transfer the denatured DNA to the membrane. Traditionally, a nitrocellulose membrane is used, although nylon or a positively charged nylon membrane may be used. Nitrocellulose typically has a binding capacity of about 100µg/cm, while nylon has a binding capacity of about 500 µg/cm. Many scientists feel nylon is better since it binds more and is less fragile. Transfer is usually done by capillary action, which takes several hours. Capillary action transfer draws the buffer up by capillary action through the gel and into the membrane, which will bind ssDNA. The Southern blot apparatus is illustrated in Figure 7.2. � Fill the container with 20x SSC so the level is just below the

container edge. � Place the gel onto the filter paper such that the open wells

are face down and the "back" of the gel is up. Make sure no bubbles are trapped between the gel and filters.

� Cut a piece of nitrocellulose to the exact size of the gel, soak the filter in 6X SSC until it is fully wet.

� Place nitrocellulose on top of the gel - again take great care to insure that no bubbles are trapped between gel and nitrocellulose.

� Cut 4-6 pieces of Whatman 3 mm paper to the same size as gel and nitrocellulose.

� Soak quickly in 6x SSC. Place these on top of nitrocellulose, again watching for trapped bubbles.

� Cut paper towels to same size as gel and filters - place these on top of the stack.

� Top with a weight. � Allow the blot to proceed, changing paper towels when the stack becomes

wet. Approximate blotting times is 14-16 hours (overnight ) for large DNA

Taking the blot apart � Peel off paper towels and Whatman 3 mm filters, trying not to dislodge the

nitrocellulose from the gel. � Flip the nitrocellulose/gel over and mark the positions of the gel wells onto

the nitrocellulose using a ball-point pen along the lower edge of the well. After marking wells, peel the gel off and discard.

� Rinse the filter in 6xSSC and air dry completely (~60 minutes at RT) Blot Fixation � After you transfer your DNA to the membrane, treat it with UV light. This

cross links (via covalent bonds) the DNA to the membrane. (You can also

bake nitrocellulose at about 80oC for a couple of hours, but be aware that it is very combustible.)

Note You may use a vacuum blot apparatus instead of capillary action. In this procedure, a vacuum sucks SSC through the membrane. This works similarly to capillary action, except more SSC goes through the gel and membrane, so it is faster (about an hour).



Buffer

Figure 7.2. Setup of the Southern blotting apparatus.

Probe the membrane with labeled ssDNA (hybridization) This process relies on the ssDNA hybridizing (annealing) to the DNA on the membrane due to the binding of complementary strands. A probe is a fragment of DNA of variable length (usually 100-1000 bases long), which is used to a nucleotide sequences that are complementary to the sequence in the probe. Must be labeled to be visualized, usually prepared by making a radioactive copy of a DNA fragment. Probing is often done with 32P labeled ATP, biotin/streptavidin or a bioluminescent probe. � To hybridize, use the same buffer as for prehybridization, but add your

specific probe.

Detection � Visualize your labeled target sequence. If radiolabeled 32P probe is used,

then you would visualize by autoradiography. Biotin/streptavidin detection is done by colorimetric methods, and bioluminescent visualization uses luminesence.


Exercise 8: Plasmid DNA Isolation Objectives � Extraction of plasmid DNA from E. Coli � Analysis of plasmid DNA by agarose gel electrophoresis and

spectrophotometer

Introduction Many types of bacteria contain plasmid DNA. Plasmids are extra-chromosomal, double-stranded circular DNA molecules generally containing 1,000 to 100,000 base pairs (Figure 8.1). Even the largest plasmids are considerably smaller than the chromosomal DNA of the bacterium, which can contain several million base pairs. Certain plasmids replicate independently of the chromosomal DNA and can be present in hundreds of copies per cell. A wide variety of genes have been discovered in plasmids. Some of them code for antibiotic resistance and restriction enzymes. Plasmids are extremely important tools in molecular cloning because they are useful in propagating foreign genes. When plasmids are used for these purposes, they are referred to as vectors.

Bacteria can sometimes take up DNA from the external environment, a natural process that we call "transformation", and sometimes the DNA taken up is a plasmid that can be maintained in the cell because it has an origin of replication. Plasmids often make the bacteria gain a gene that gives a selective advantage to the cell, which then replicates (instead of dying) and makes more copies of the plasmid. The plasmids used in transformation typically have three important elements:

• A cloning site (a place to insert foreign DNAs) • An origin of replication • A selectable marker gene (e.g. resistance to

ampicillin)

Plasmids can be classified into 5 classes according to their function:

1. Fertility-F-plasmids, Facilitate bacterial conjugation.

2. Resistance-(R)plasmids, which contain genes that can build a resistance against antibiotics or poisons.

3. Col-plasmids, which contain genes that code for bacteriocins, proteins that can kill other bacteria.

4. Degradative plasmids, which enable the digestion of unusual substances, e.g., toluene or salicylic acid.

5. Virulence plasmids, which turn the bacterium into a pathogen.

Figure 8.1. "A" represents a bacterial cell containing plasmids. "B" represents electron micrograph of Plasmids.

8

Exercixe 8 Plasmid DNA Isolation


Isolation of plasmid DNA from bacterial cells is an essential step for many molecular biology procedures. Many protocols for large- and small-scale isolation of plasmids (mini-preps) have been published. The plasmid purification procedures, unlike the procedures for purification of genomic DNA, should involve removal not only of protein, but also another major impurity: bacterial chromosomal DNA. To achieve separation of plasmid from chromosomal DNA, these methods exploit the structural differences between plasmid and chromosomal DNA. Plasmids are circular supercoiled DNA molecules substantially smaller than bacterial chromosomal DNA. Plasmid DNA isolation involves (1) growth of bacteria with amplification of plasmid, (2) harvesting and lysis of bacteria, and (3) purification of plasmid DNA. Normally there are ten to two hundred plasmids (relaxed - not connected to chromosomal replication) per bacterial cell. The bacteria containing the plasmid grow in LB medium plus antibiotic that selects for plasmid (antibiotic resistant gene located on plasmid). By gentle bacterial lysis small molecules, including covalently closed supercoiled plasmids are released into solution. Mild alkali treatment to break most of the hydrogen bonds in DNA and degrade chromosomal DNA. Closed circular plasmids regain their native configuration when returned to neutral pH while larger linear chromosomal DNA fragments remain in the denature state trapped in the cell debris. The cell debris is precipitated and the supernatant containing the plasmid is collected and plasmid is isolated. The yield of plasmid DNA is dependent on the plasmid copy number, plasmid type, bacterial strain, and growth conditions. Procedure Materials

� LB (Luria-Bertani) containing 20 mg/l ampicilin � GTE buffer (50mM Glucose, 25 mM Tris-Cl & 10mM EDTA, pH 8) � NaOH/ SDS lysis solution (0.2 M NaOH, 1% SDS) � 5 M potassium acetate solution (pH 4.8) � Isopropanol

� TE buffer � RNAse solution (20 mg/ml stock) � PCIA (phenol/chloroform/isoamyl alcohol) � 7.5 M ammonium acetate solution

� Absolute ethanol DNA Plasmid Miniprep Protocol

1. Pick single colony and inoculate 5 ml of LB broth containing 20 mg/l ampicillin or 0.1mg/5ml. Optional: Use a 15ml conical tube with a loosened cap and a piece of tape to hold it in place. Shake at 250 RPM 37oC overnight.

2. Centrifuge 1.5mL cells in 1.5 mL Eppendorf tube at top speed for 1 minute. Aspirate supernatant.

Exercixe 8 Plasmid DNA Isolation


3. Resuspend cell pellet in 100 ul of GTE buffer. Vortex gently if necessary.

4. Add 200 µl of NaOH/SDS lysis solution. Invert tube 6-8 times.

5. IMMEDIATELY add 150 µl of 5 M potassium acetate solution (pH 4.8). This solution neutralizes NaOH in the previous lysis step while precipitating the genomic DNA and SDS in an insoluble white, rubbery precipitate. Spin at top speed 1 min.

6. Transfer supernatant to a new tube, being careful not to pick up any white flakes. Precipitate the nucleic acids with 0.5mL of isopropanol on ice for 10 minutes and centrifuge at top speed for 1 minute.

7. Aspirate off all the isopropanol supernatant. Dissolve the pellet in 0.4 ml of TE buffer. Add 10µl of RNAse solution (20 mg/ml stock stored at -20 °C), vortex and incubate at 37°C for 20 to 30 minutes to digest remaining RNA.

8. Extract proteins from the plasmid DNA using PCIA (phenol/ chloroform/isoamyl alcohol) by adding about 0.3 ml. Vortex vigorously for 30 seconds. Centrifuge at full speed for 5 minutes at room temperature. Note organic PCIA layer will be at the bottom of the tube.

9. Remove upper aqueous layer containing the plasmid DNA carefully avoiding the white precipitated protein layer above the PCIA layer, transferring to a clean 1.5 ml eppendorf tube.

10. Add 100 ml of 7.5 M ammonium acetate solution and 1 ml of absolute ethanol to precipitate the plasmid DNA on ice for 10 minutes. Centrifuge at full speed for 5 minutes at room temperature.

11. Aspirate off ethanol solution and resuspend or dissolve plasmid DNA pellet in 50µl of DNA buffer.

12. Prepare a 1% agarose gel, load the plasmid DNA sample after mixing with loading buffer and run the gel for 30 minutes.

13. Measure the concentration of the plasmid by a spectrophotometer.

14. The purity of the isolated plasmid DNA can also be measured by spectrophotometer. DNA UV absorbance peaks at 260 nm, while protein UV absorbance peaks at 280 nm. The ratio of the absorbance at 260 nm/280 nm is a measure of the purity of a DNA sample from protein contamination; it should be between 1.7 and 2.0.

The ratio of the absorbance at 260 nm/230 nm is a measure of the purity of a DNA sample from organics and/or salts; it should be about 2.0. Low 260/230 ratio indicates contamination by organics and/or salts.


Exercise 9: Transformation of Escherichia coli Objectives: � Understand the transformation procedure using the heat shock method. � Understand how DNA can be transferred to an organism and the change

in phenotype that may result from such a transfer. � Transformation of a gene for resistance to the antibiotic ampicillin into a

bacterial strain (E. coli) that is sensitive to ampicillin.

Introduction Transformation is a very basic technique that is used on a daily basis in a molecular biological laboratory. The purpose of this technique is to introduce a foreign plasmid into a bacteria and to use that bacteria to amplify the plasmid in order to make large quantities of it. This is based on the natural function of a plasmid, i.e. transfer genetic information vital to the survival of the bacteria. In molecular biology, transformation refers to a form of genetic exchange in which the genetic material carried by an individual cell is altered by incorporation of foreign (exogenous) DNA. This foreign DNA may be derived from unrelated species and even other kingdoms, such as bacteria, fungi, plants or animals. Bacteria and yeast have been transformed with human genes to produce proteins that are useful in treating human diseases and disorders e.g. the production of insulin. Some bacteria have been modified such that they are able to digest oil from accidental spills. Transformation is usually more difficult with multicellular organisms such as plants, in which it is necessary to either alter many cells with the new piece of DNA or to alter just a single cell and then induce it to produce a whole, new plant. Genetic transformation of plants and other organisms does occur naturally. The bacterium you will be transforming, Escherichia coli (E.coli), Figure 9.1, lives in the human gut and is a relatively simple and well understood organism. Its genetic material consists mostly of one large circle of DNA 3-5 million base pairs in length, with small loops of DNA called plasmids, usually ranging from 5,000-10,000 base pairs in length, present in the cytoplasm. It is these plasmids that bacteria can transfer back and forth, allowing them to share genes among one another and thus to naturally adapt to new environments. The ability of bacteria to maintain these plasmids and replicate them during normal cell multiplication is the basis of cell transformation. The plasmids are used as “gene taxis” in transformation events to bring DNA of interest into the cell where it can integrate into the genome (or remain as a plasmid within a bacterium) and be translated into proteins not normally found in that organism. The plasmid which is going to be used in this experiment contains an ampicillin-resistance gene. Ampicillin is an antibiotic and works by preventing

Figure 9.1. Electron micrograph of a cluster of E. coli magnified 10,000 times.

9

Exercixe 9 Transformation of Escherichia coli


E.coli from constructing cell walls, thereby killing the bacteria. When the ampicillin-resistance gene is present it directs the production of an enzyme that blocks the action of the ampicillin and the bacteria are able to survive. Bacteria without the plasmid and hence the resistance gene are unable to grow on a plate containing ampicillin in the medium and only the transformants will survive. Bacterial strains may have natural competence, i.e. they have the ability to take up DNA from the medium. Natural competence is a genetically programmed physiological state. Natural transformation is distinct from artificial transformation by techniques such as electroporation. In addition, some bacterial strains, such as E. coli, can be made artificially competent using CaCl2 and heat shock treatment. Since DNA is a very hydrophilic molecule, it won't normally pass through a bacterial cell's membrane. In order to make bacteria take in the plasmid, they must first be made "competent" to take up DNA. This is done by creating small holes in the bacterial cells by suspending them in a solution with a high concentration of calcium. DNA can then be forced into the cells by incubating the cells and the DNA together on ice, placing them briefly at 42oC (heat shock), and then putting them back on ice. This causes the bacteria to take in the DNA. The cells are then plated out on antibiotic containing media. The procedure to prepare competent cells can sometimes be tricky. Bacteria aren't very stable when they have holes put in them, and they die easily. A poorly performed procedure can result in cells that aren't very competent to take up DNA. A well- performed procedure will result in very competent cells. The competency of a stock of competent cells is determined by calculating how many E. coli colonies are produced per microgram (10 -6 grams) of DNA added. An excellent preparation of competent cells will give ~108 colonies per µg. A poor preparation will be about 10 4 / µg or less. Our preps should be in the range of 10 5 to 10 6.

Procedure In this experiment you will be making competent cells, transforming them with a plasmid and calculating their competency.

Materials

� Luria-Bertani media � Luria-Bertani broth � 50mM CaCl2 solution � Ampicillin (10 mg/ml) � Ampicillin resistant Plasmid (0.005 µg/µl)



Method

Competency � Label two sterile microtubes: one “+”and the other “-” � Using a disposable pipette, add 250 µl of 50mM CaCl2 solution to each

tube (“+” and “-”) and place them both on ice � Use a sterile plastic loop to transfer one or two 3 mm bacterial colonies

to the “+” tube, return tube to ice (Do not pick up any agar as it may inhibit the transformation process)

� Transfer a mass of cells to the “-” tube � Add 10 µl of ampicillin resistant plasmid directly into the

CaCl2 in “+” tube

� Return the “+” tube to the ice. DO NOT add the plasmid to your "-" tube. Incubate both tubes on ice for 15 minutes

Plasmid Insertion � Remove the tubes from the ice and immediately hold them in a 42oC

water bath for 90 seconds. (The marked temperature change causes the cells to readily absorb the plasmid DNA)

� Move test tubes suddenly from the water bath back to the ice � Keep on ice for at least 1 minute

Recovery period � Allow the cell to regain strength and start to multiply � Add 250 µl sterile Luria Broth to both tubes using a sterile pipette � Move tubes to water bath at 37oC for 5 min

Growth & Isolation � Label four LB media plates as follows:

o Plate number 1, LB -no plasmid o Plate number 2, LB/ Amp – no plasmid o Plate number 3, LB + plasmid o Plate number 4, LB/ Amp + no plasmid

� Transfer 100 µl of cell suspension to each plate by using sterile transfer

pipette � Lift the lid of the plates and sweep the drop of cell suspension with a loop

to distribute it over the surface of the plate � Repeat this for the remaining 3 plates � Incubate the plates upside down for 24 hours at 37oC. � Analyze the results of the transformation

Determination of Transformation Efficiency

Transformation efficiency is a quantitative determination of how many cells were transformed per 1 µg of plasmid DNA. In essence, it is an indicator of how well the transformation experiment worked. You will calculate the transformation efficiency from the data you collect from your experiment.

Note The cells are kept cold to prevent them from growing while the plasmids are being absorbed.



� Count the number of colonies on the plate with ampicillin that is labeled LB / Amp +

� A convenient method to keep track of counted colonies is to mark the colony with a lab marking pen on the outside of the plate

Determine the transformation efficiency using the formula:

final vol at recovery (ml) Number of transformants Number of transformants per µg

= vol plated (ml)

X µg of DNA


Exercise 10: SDS-PAGE Objectives � To understand the principle of Sodium DodecylSulphate- PolyAcrylamide

Gel Electrophoresis (SDS-PAGE) � To become familiar with the SDS-PAGE setup

Introduction A Polyacrylamide Gel is a separation matrix used in electrophoresis of biomolecules, such as proteins or DNA fragments. Scientists used polyacrylamide gels to separate DNA fragments differing by a single base-pair in length. Most modern DNA separation methods now use agarose gels, except for particularly small DNA fragments. It is currently most often used in the field of immunology and protein analysis, often used to separate different proteins or isomers of the same protein into separate bands. These can be transferred onto a nitrocellulose to be probed with antibodies and corresponding markers, such as in a western blot. Proteins are a highly diversified class of biomolecules. Differences in their chemical properties, such as charge, functional groups, shape, size and solubility enable them to perform many biological functions. Determination of the molecular weight of a protein is of fundamental importance to its biochemical characterization. If the amino acid composition or sequence is known, the exact molecular weight of a polypeptide can be calculated. SDS gel electrophoresis is commonly used to obtain reliable molecular weight estimates for denatured polypeptides. A protein can have a net negative or net positive charge, depending on its amino acid composition and the pH. At certain pH values of solutions, the molecule can be electrically neutral, i.e. negative and positive charges are balanced. In this case, the protein is isoelectric. In the presence of an electrical field, proteins with net charges will migrate towards the electrodes of opposite charge. Proteins exhibit different three-dimensional shapes and folding patterns which are determined by their amino acid sequences and intracellular processing. The physical-chemical properties of proteins affect the way they migrate during gel electrophoresis. Gels used in electrophoresis (e.g. agarose, polyacrylamide) consist of microscopic pores of a defined size range that act as a molecular sieve. Only molecules with net charge will migrate through the gel when it is in an electric field. Small molecules pass through the pores more easily than large ones. Molecules having more charge than others of the same shape and size will migrate faster. Molecules of the same mass and charge can have different shapes. In such cases, those with more compact shape (sphere-like) will migrate through the gel more rapidly than those with an elongated shape, like a rod. In summary, the charge, size and shape of a native protein all affect its electrophoretic migration rates. Electrophoresis of native proteins is useful in the clinical and immunological analysis of complex

10

Exercixe 10 SDS-PAGE


biological samples, such as serum, but is not reliable to estimate molecular weights.

Polyacrylamide gel electrophoresis Polyacrylamide gels are formed by mixing the monomer, acrylamide, the cross-linking agent, methylenebisacrylamide, free radical generator, ammonium persulfate, and tetramethylethylenediamine (TEMED) (TEMED accelerates the rate of formation of free radicals from persulfate) in aqueous buffer. Free radical polymerization of the acrylamide occurs. At various points the acrylamide polymers are bridged to each other (Figure 10.1). The pore size in polyacrylamide gels is controlled by the gel concentration and the degree of polymer cross-linking. The electrophoretic mobility of the proteins is affected by the gel concentration. Higher percentage gels are more suitable for the separation of smaller polypeptides. The polymerization process is inhibited by oxygen. Consequently, polyacr-ylamide gels are most often prepared between glass plates separated by strips called spacers. As the liquid acrylamide mixture is poured between the plates, air is displaced and polymerization proceeds more rapidly. Sodium dodecylsulfate (SDS) binds strongly to most proteins and causes them to unfold to a random, rod-like chains (Figure 10.2). No covalent bonds are broken in this process. The end result has two important features: 1) all proteins contain only primary structure and 2) all proteins have a large negative charge (Figure 10.2), which means they will all migrate towards the positive pole when placed in an electric field. Proteins which contain several polypeptide chains that are associated only by noncovalent forces will be dissociated by SDS into separate, denatured polypeptide chains. Proteins can contain covalent crosslinks known as disulfide bonds. High concentrations of reducing agents, such as β-mercaptoethanol, can break disulfide bonds.

Caution

It should be noted that acrylamide is a neuron-toxin and can be absorbed through the skin.

Figure 10.1. Schematic diagram of polyacrylamide polymer formation.

Figure 10.2. A protein before adding SDS and after adding SDS.



The amount of negative charge of the SDS is much more than the negative and positive charges of the amino acid residues. The large quantity of bound SDS efficiently masks the intrinsic changes in the protein. Consequently, SDS denatured proteins are net negative and since the binding of the detergent is proportional to the mass of the protein, the charge to mass ratio is constant. The shape of SDS denatured proteins are all rod-like.

During SDS electrophoresis, the proteins migrate through the gel towards the positive electrode at a rate that is inversely proportional to their molecular weight. In other words, the smaller the denatured polypeptide, the faster it migrates. The molecular weight of an unknown polypeptide is obtained by the comparison of its position after electrophoresis to the positions of standard SDS denatured proteins. The molecular weights of the standard proteins have been previously determined. After proteins are visualized by staining and destaining, their migration distance is measured. The log10 of the molecular

weights of the standard proteins are plotted versus their migration distance. The molecular weight of unknowns are then easily calculated from the standard curve. Procedure Materials

� Acrylamide/ Bisacrylamide (40%) � Tris-HCl (1 M – pH 8.8) � Tris-HCl (0.5 M – pH 6.8) � Distilled water � 10% SDS � 10% Ammonium Persulfate � TEMED � Sample buffer � Silver stain kit

Methods � Label 2 beakers one "running" and the other "stacking" � The gel is prepared by adding the following components to the

corresponding beaker

Reagent 8% (Running Gel) 5% (Stacking Gel) Acrylamide/ Bisacrylamide (40%) * 4.0 mls 2.5 mls

Tris-HCl 7.5 mls

(1 M – pH 8.8) 7.5 mls

(0.5 M – pH 6.8) water (distilled) 8.2 mls 9.7 mls 10% SDS 200 µl 200 µl 10% Ammonium Persulfate 100 µl 100 µl TEMED (added last) 10 µl 10 µl

* = 19:1 w:w ratio of acrylamide to N,N'-methylene bis-acrylamide � Mix ingredients gently in the order shown above, ensuring no air bubbles

form � Assemble two glass plates with two side spacers � Pour into glass plate assembly, first the running gel and then the

stacking gel



� Insert comb, allow to set, and then remove comb after gel has set (15 min)

� Assemble glass plates onto the electrophoresis unit, fill with electrophoresis buffer

� Dilute samples at least 1:4 with sample buffer, heat at 95oC for 4 minutes prior to loading

� Load the samples onto polyacrylamide gel � Run at 200 volts for 30-40 minutes � When the tracking dye reaches the bottom of the gel, turn off the power

supply � Pour out the buffer by inverting the entire unit over a sink � Open the gel sandwich. Remove the spacers and peel the gel off the

plate into a tray of stain. Wetting the gel helps to loosen it from the plastic plate

� Stain with Silver stain (follow the instructions of the kit)

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