Molecular Biology Finale

(An Exclusive Biotechnology Institute)

Submitted by:

Nitin Saxena

B.tech biotech

4th semester

Jaipur National University

This is to acknowledge the valuable contribution of the following individuals:

I feel privileged to express my sense of gratitude to Ms. Neha Sharma, faculty Incharge, Molecular Biology, Dr. B. Lal institute, Jaipur. For allowing me to work and for her great guidance and support on every step, I required.

I would like to thank Mr. Deepesh, Lab Assistant, Dr. B. Lal institute, Jaipur for his valuable help.

A sincere vote of thanks to my batch mates also for making this one month training more of fun than just a mere part of study curriculum.

Contents:

1. Introduction to Molecular Biology 1.1. About 1.2. Historical background 1.3. Techniques associated cum applications

2. Instrumentation 2.1. Laminar air flow and biosafety chamber 2.2. Centrifuge 2.3. UV – visible spectrophotometer 2.4. Electrophoresis unit 2.5. Thermal cycler 2.6. Autoclave 2.7. Incubator 2.8. Dry bath 2.9. UV transilluminator 2.10. Vortex 2.11. Hot plate 2.12. pH meter 2.13. Weighing balance

3. Module exercise

3.1. Preparation of buffer and reagents

3.2. Isolation of Genomic DNA from Prokaryotes

3.3. Isolation of Genomic DNA from Plants

3.4. Qualitative and Quantitative analysis of DNA

3.5. Determination of melting temperature of DNA

3.6. SDS PAGE

3.7. PCR technique

1.1. About

Molecular biology is the study of biologybiology and chemistry, particularly with understanding the interactions between the various systems of abetween DNA, RNA and protein biosynthesis

According to William Astbury, as per his writing in Nature in 1961;

“Molecular Biology is not much a technique but an approach,called basic sciences with the leading idea of searching below the largebiology for the corresponding molecular plan. It is concerned particularly with theand is predominantly three-dimensional and structuralrefinement of morphology. It must at the same t

Schematic relationship between biochemistry, genetics, and molecular biology

1.2. Historical background

While molecular biology was established in the 1930s, the term 1938. Warren was the director of Natural Sciences for thebelieved that biology was about to undergo a period of significant change given recent advances in fields such as X-ray crystallography. He therefore channeled significant amounts of (Rockefeller Institute) money into biological fields.

biology at a molecular level. This field overlaps with other areas of genetics and biochemistry. Molecular biology chiefly concerns itself

with understanding the interactions between the various systems of a cell, including the interactions protein biosynthesis as well as learning how these interactions are regulated.

as per his writing in Nature in 1961;

iology is not much a technique but an approach, an approach from the viewpoint of the socalled basic sciences with the leading idea of searching below the large-scale manifestations of classical

or the corresponding molecular plan. It is concerned particularly with the forms of biological molecules dimensional and structural—which does not mean, however, that it is merely a

refinement of morphology. It must at the same time inquire into genesis and function.”

Schematic relationship between biochemistry, genetics, and molecular biology

Historical background

While molecular biology was established in the 1930s, the term was first coined by Warren Weaver1938. Warren was the director of Natural Sciences for the Rockefeller Foundationbelieved that biology was about to undergo a period of significant change given recent advances in fields

. He therefore channeled significant amounts of (Rockefeller Institute)

level. This field overlaps with other areas of . Molecular biology chiefly concerns itself

, including the interactions as well as learning how these interactions are regulated.

an approach from the viewpoint of the so-scale manifestations of classical

of biological molecules which does not mean, however, that it is merely a

Warren Weaver in Rockefeller Foundation at the time and

believed that biology was about to undergo a period of significant change given recent advances in fields . He therefore channeled significant amounts of (Rockefeller Institute)

1.3. Techniques associated

Since the late 1950s and early 1960s, molecular biologists have learned to characterize, isolate, and manipulate the molecular components of cells and organisms. These components include DNA, the repository of genetic information; RNA, a close relative of DNA whose functions range from serving as a temporary working copy of DNA to actual structural and enzymatic functions as well as a functional and structural part of the translational apparatus; and proteins, the major structural and enzymatic type of molecule in cells.

Expression cloning One of the most basic techniques of molecular biology to study protein function is expression cloning. In this technique, DNA coding for a protein of interest is cloned (using PCR and/or restriction enzymes) into a plasmid (known as an expression vector).

Polymerase chain reaction (PCR) The polymerase chain reaction is an extremely versatile technique for copying DNA. In brief, PCR allows a single DNA sequence to be copied (millions of times), or altered in predetermined ways.

Gel electrophoresis Gel electrophoresis is one of the principal tools of molecular biology. The basic principle is that DNA, RNA, and proteins can all be separated by means of an electric field. In agarose gel electrophoresis, DNA and RNA can be separated on the basis of size by running the DNA through an agarose gel. Proteins can be separated on the basis of size by using an SDS-PAGE gel, or on the basis of size and their electric charge by using what is known as a 2D gel electrophoresis.

Macromolecule blotting and probing

a. Southern blotting Named after its inventor, biologist Edwin Southern, the Southern blot is a method for probing for the presence of a specific DNA sequence within a DNA sample. DNA samples before or after restriction enzyme digestion are separated by gel electrophoresis and then transferred to a membrane by blotting via capillary action. The membrane is then exposed to a labeled DNA probe that has a complement base sequence to the sequence on the DNA of interest. Most original protocols used radioactive labels, however non-radioactive alternatives are now available.

b. Northern blotting

The northern blot is used to study the expression patterns of a specific type of RNA molecule as

relative comparison among a set of different samples of RNA. It is essentially a combination

of denaturing RNA gel electrophoresis, and a blot. In this process RNA is separated based on size and

is then transferred to a membrane that is then probed with a labeled complement of a sequence of

interest. The results may be visualized through a variety of ways depending on the label used;

however, most result in the revelation of bands representing the sizes of the RNA detected in sample.

The intensity of these bands is related to the amount of the target RNA in the samples analyzed. The

procedure is commonly used to study when and how much gene expression is occurring by measuring

how much of that RNA is present in different samples. It is one of the most basic tools for determining

at what time, and under what conditions, certain genes are expressed in living tissues.

c. Western blotting

Antibodies to most proteins can be created by injecting small amounts of the protein into an animal

such as a mouse, rabbit, sheep, or donkey (polyclonal antibodies) or produced in cell culture

(monoclonal antibodies). These antibodies can be used for a variety of analytical and preparative

techniques.

In western blotting, proteins are first separated by size, in a thin gel sandwiched between two glass

plates in a technique known as SDS-PAGE (sodium dodecyl sulfate polyacrylamide gel

electrophoresis). The proteins in the gel are then transferred to a PVDF, nitrocellulose, nylon or other

support membrane. This membrane can then be probed with solutions of antibodies. Antibodies that

specifically bind to the protein of interest can then be visualized by a variety of techniques, including

colored products, chemiluminescence, or autoradiography. Often, the antibodies are labeled with an

enzyme. When a chemiluminescent substrate is exposed to the enzyme it allows detection. Using

western blotting techniques allows not only detection but also quantitative analysis.

Analogous methods to western blotting can be used to directly stain specific proteins in

live cells or tissue sections. However, these immunostaining methods, such as FISH, are used more

often in cell biology research.

d. Eastern blotting

Eastern blotting technique is to detect post-translational modification of proteins. Proteins blotted on to

the PVDF or nitrocellulose membrane are probed for modifications using specific substrates.

The terms northern, western and eastern blotting are derived from what initially was a molecular biology joke

that played on the term Southern blotting, after the technique described by Edwin Southern for the hybridization

of blotted DNA. Patricia Thomas, developer of the RNA blot which then became known as the northern

blot actually didn't use the term Further combinations of these techniques produced such terms

as southwestern (protein-DNA hybridizations), northwestern (to detect protein-RNA interactions) and far

westerns (protein-protein interactions), all of which are presently found in the literature.

Arrays

A DNA array is a collection of spots attached to a solid support such as a microscope slide where each spot

contains one or more single-stranded DNA oligonucleotide fragment. Arrays make it possible to put down a

large quantity of very small (100 micrometer diameter) spots on a single slide.

Arrays can also be made with molecules other than DNA. For example, an antibody array can be used to

determine what proteins or bacteria are present in a blood sample.

Allele Specific Oligonucleotide

Allele specific oligonucleotide (ASO) is a technique that allows detection of single base mutations without the

need for PCR or gel electrophoresis.

The Illumina Methylation Assay is an example of a method that takes advantage of the ASO technique to measure one base pair differences in sequence.

Antiquated technologies In molecular biology, procedures and technologies are continually being developed and older technologies abandoned. For example, before the advent of DNA gel electrophoresis (agarose or polyacrylamide), the size of DNA molecules was typically determined by rate sedimentation in sucrose gradients, a slow and labor-intensive technique requiring expensive instrumentation; prior to sucrose gradients,viscometry was used.

Aside from their historical interest, it is often worth knowing about older technology, as it is occasionally useful to solve another new problem for which the newer technique is inappropriate.

2.1. Laminar flow cabinet

A laminar flow cabinet or laminar flow closet or tissue culture hood is a carefully enclosed bench designed to prevent contamination of semiconductor wafers, biological samples, or any particle sensitive device. Air is drawn through a HEPA filter and blown in a very smooth, laminar flow towards the user. The cabinet is usually made of stainless steel with no gaps or joints where spores might collect.

Such hoods exist in both horizontal and vertical configurations, and there are many different types of cabinets with a variety of airflow patterns and acceptable uses.

Working principle

An air stream, sterilized by an H14 filter is directed as laminar air flow towards the working surface ensuring total particle elimination in the work space, the air is exhausted trough the frontal access, avoiding any contamination. Laminar Air Flow is based on the flow of air current to create uniform velocity, along parallel lines, which helps in transforming microbial culture in aseptic conditions .

Benefits of laminar flow cabinet:

1. Provide clean air to the working area. 2. Provide a constant flow of air out of the work area to prevent room air from entering. 3. The air flowing out from the hood suspends and removes contaminants introduced into the work area by personnel. The most important part of a laminar flow hood is a high efficiency bacteria-retentive filter. Room air is taken into the unit and passed through a pre-filter to remove gross contaminants (lint, dust etc). The air is then compressed and channeled up behind and through the HEPA filter (High Efficiency Particulate Air filter) in a laminar flow fashion--that is the purified air flows out over the entire work surface in parallel lines at a uniform velocity. The HEPA filter removes nearly all of the bacteria from the air. Why control room air? The environmental control of air is of concern because room air may be highly contaminated. Example: Sneezing produces 100,000 - 200,000 aerosol droplets which can then attach to dust particles. These contaminated particles may be present in the air for weeks.

Points to keep in mind

1. A direct path must be maintained between the filter and the area inside the hood where the manipulations are being performed. Air downstream from non-sterile objects (such as solution containers, hands etc.) becomes contaminated from particles blown off these objects. To best illustrate this very important point click on the two examples of proper aseptic technique: (manipulation of a vial and an ampoule).

2. Always minimize clutter. Waste and other items should never enter the hood. All calculations should be done before entering the hood.

3. 3. Wash hands and arms before compounding or re-entering the hood. Also, remove any jewelry from the hands and wrists. It is important that you keep your hands within the cleaned area of the hood as much as possible. Do not touch your hair, face or clothing.

4. Excess dust should be removed from items before introducing them into the hood.

5. Arrange objects in a manner to get full benefit of the laminar flow of air. Critical items should be placed as close to the air source as possible. In a horizontal hood, items should be placed no closer than 3 inches from the very back of the hood (nothing should touch the filter).

6. Occasionally, you may stack a few items (eg IVPB's), however they must be stacked from lower to higher starting from the back of the hood. Also, it should be limited to a maximum of 3 to 4 items.

7. When working in a horizontal laminar flow hood, all work must be performed at a distance of no less than 6 inches from the front edge of the work surface. At a distance of less than 6 inches, laminar flow air begins to mix with the outside air and contamination is possible. Never become so engrossed in your work that you forget this basic rule.

8. Avoid spraying or squirting solutions onto the HEPA filter. Always aim away from the filter when opening ampoules or adjusting syringes.

9. Outer pouches and wraps should be removed at the edge of the work area as the sterile contents are pulled into the work area. Never bring these items into the main work area.

10. Large objects should never be placed near the back of the hood. Not only do these objects contaminate everything downstream, but they also disrupt the laminar flow pattern of air which normally suspends the contaminants and removes them from the area.

11. Remember that hand cleanliness is further reduced each time more bottles and other non-sterile items are handled.

12. Anytime something is spilled, the work surface of the laminar flow hood should be thoroughly cleaned with alcohol. A long side to side motion should be used starting at the back of the hood and then working forward. The acrylic plastic sides should also be cleaned periodically.

13. It is possible to overcome the established airflow velocity by a strong reverse current produced by coughing, quick movements, talking etc. Keep all of these to a minimum in order to maintain a sterile environment. Do not talk, cough or sneeze into the hood!

Standard operation procedure of laminar air flow

1. Switch on the electric supply 2. Switch on the florescent lamp and blower 3. Open the front panel of lf 4. Prior to start clean the lf bench with bench cleaning/ disinfectant solution like 75% ethanol 5. Cover the front panel of lf and switch off the florescent lamp and blower 6. Switch on the UV lamp for 15-20 minutes and ensure to avoid the direct exposure to UV rays as

they are harmful and are mutagenic 7. Switch off the UV lamp after desired time 8. Switch on the florescent lamp and blower 9. Open the front panel of lf 10. Turn the flame on for maintaining aseptic condition 11. Ensure safety wears like mask , gloves, apron during work

12. While working ; the operator should have well trimmed nail and the hair should be tied properly 13. Care should be taken to keep the flame and disinfectant separately 14. Any spillage should be immediately cleaned with the alcohol swab 15. After completion of work clear the lf bench with disinfectant 16. Close the front panel of lf 17. Switch on the UV lamp for 5-10 minutes and ensure to avoid the direct exposure to UV rays as

they are harmful and are mutagenic 18. Switch off the UV lamp after desired time

2.2. Centrifuge A centrifuge is a piece of equipment, generally driven by an electric motor (some older models were spun by hand), that puts an object in rotation around a fixed axis, applying a force perpendicular to the axis. The centrifuge works using the sedimentation principle, where the centripetal acceleration causes more dense substances to separate out along the radial direction (the bottom of the tube). By the same token, lighter objects will tend to move to the top (of the tube; in the rotating picture, move to the centre).

In the picture shown, the rotating unit, called the rotor, has fixed holes drilled at an angle (to the vertical). Test tubes are placed in these slots and the rotor is spun. As the centrifugal force is in the horizontal plane and the tubes are fixed at an angle, the particles have to travel only a little distance before they hit the wall and drop down to the bottom. These angle rotors are very popular in the lab for routine use.

Working principle Protocols for centrifugation typically specify the amount of

rather than specifying a rotational speed

quoted in multiples of g, the standard acceleration due to

is important because two rotors with different diameters running at the same rotational speed will

subject samples to different accelerations.

Since the motion is circular the acceleration can be calculated as the product of the

square of the angular velocity. Traditionally named "relative centrifugal force" (RCF), it is the

measurement of the acceleration applied to a sample within a centrifuge and it is measured i

gravity (times gravity or × "g"). It is given by

where

is earth's gravitational acceleration

is the rotational radius,

is the rotational speed, measured in revolutions per unit of time.

When the rotational speed is given in

expressed in centimetres (cm) the above relationship becomes

where

is the rotational radius measured in

is rotational speed measured in

Standard operation procedure of centrifuge

1. Switch on the electric supply2. Place the sample vials 3. Always use counter balance4. Set the required time duration in minutes by “set time” key5. Set the required rpm by rotating the knob6. Use “break” button if in between break is required7. After centrifuge remove the vials8. Switch off the machine

Protocols for centrifugation typically specify the amount of acceleration to be applied to the sample,

rotational speed such as revolutions per minute. The acceleration is often

g, the standard acceleration due to gravity at the Earth's surface. This distinction

is important because two rotors with different diameters running at the same rotational speed will

subject samples to different accelerations.

the acceleration can be calculated as the product of the radius

. Traditionally named "relative centrifugal force" (RCF), it is the

measurement of the acceleration applied to a sample within a centrifuge and it is measured i

"g"). It is given by

gravitational acceleration,

, measured in revolutions per unit of time.

When the rotational speed is given in revolutions per minute (RPM) and the rotational radius is

(cm) the above relationship becomes

is the rotational radius measured in centimetres (cm),

measured in revolutions per minute (RPM)

Standard operation procedure of centrifuge

Switch on the electric supply

Always use counter balance Set the required time duration in minutes by “set time” key

the required rpm by rotating the knob Use “break” button if in between break is required After centrifuge remove the vials

to be applied to the sample,

. The acceleration is often

the Earth's surface. This distinction

is important because two rotors with different diameters running at the same rotational speed will

radius and the

. Traditionally named "relative centrifugal force" (RCF), it is the

measurement of the acceleration applied to a sample within a centrifuge and it is measured in units of

(RPM) and the rotational radius is

Uses Isolating suspensions

Simple centrifuges are used in chemistry, biology, and biochemistry for isolating and separating

suspensions. They vary widely in speed and capacity. They usually comprise a rotor containing two, four,

six, or many more numbered wells within which the samples containing centrifuge tips may be placed.

Isotope separation

Other centrifuges, the first being the Zippe-type centrifuge, separate isotopes, and these kinds of

centrifuges are in use in nuclear power and nuclear weapon programs.

Gas centrifuges are used in uranium enrichment. The heavier isotope of uranium (uranium-238) in

the uranium hexafluoride gas tends to concentrate at the walls of the centrifuge as it spins, while the

desired uranium-235 isotope is extracted and concentrated with a scoop selectively placed inside the

centrifuge. It takes many thousands of centrifuges to enrich uranium enough for use in a nuclear

reactor (around 3.5% enrichment), and many thousands more to enrich it to weapons-grade (around

90% enrichment) for use in nuclear weapons.

Commercial applications

• Centrifuges with a batch weight of up to 2,200 kg per charge are used in the sugar industry to

separate the sugar crystals from the mother liquor.

• Standalone centrifuges for drying (hand-washed) clothes – usually with a water outlet.

• Centrifuges are used in the attraction Mission: SPACE, located at Epcot in Walt Disney World,

which propels riders using a combination of a centrifuge and a motion simulator to simulate the

feeling of going into space.

• In soil mechanics, centrifuges utilize centrifugal acceleration to match soil stresses in a scale

model to those found in reality.

• Large industrial centrifuges are commonly used in water and wastewater treatment to

dry sludges. The resulting dry product is often termed cake, and the water leaving a centrifuge

after most of the solids have been removed is called centrate.

• Large industrial centrifuges are also used in the oil industry to remove solids from the drilling

fluid.

• Disc-stack centrifuges used by some companies in Oil Sands industry to separate small amounts

of water and solids from bitumen

• Centrifuges are used to separate cream (remove fat) from milk.

2.3. Autoclave

An autoclave is a device to sterilize equipment and supplies by subjecting them to high pressure steam

at 121 °C or more, typically for 15 to 20 minutes depending on the size of the load and the contents. It

was invented by Charles Chamberland in 1879, although a precursor known as the steam digester was

created by Denis Papin in 1679. The name comes from Greek auto, ultimately meaning self, and

Latin clavis meaning key — a self-locking device.

Working principle The principle in auto clave is sterilizing the material using steam and pressure. With the increase in

pressure, temperature also increases thus raising the boiling temperature of water from 100oC to 121oC.

With high pressure, the steam so formed cause rupture of microbial cell wall and at very high

temperature and pressure the water content is absorbed and with absence of water the microbes die

thus sterilizing the material. The principle is basically same as that in an ordinary pressure cooker;

however the temperature and pressure used is higher.

Air removal It is very important to ensure that all of the trapped air is removed, as hot air is very poor at achieving

sterility. Steam at 134 °C can achieve in 3 minutes the same sterility that hot air at 160 °C takes two

hours to achieve. Methods of achieving air removal include:

Downward displacement (or gravity type) - As steam enters the chamber, it fills the upper areas as it is

less dense than air. This compresses the air to the bottom, forcing it out through a drain. Often a

temperature sensing device is placed in the drain. Only when air evacuation is complete should the

discharge stop. Flow is usually controlled through the use of a steam trap or a solenoid valve, but bleed

holes are sometimes used, often in conjunction with a solenoid valve. As the steam and air mix it is also

possible to force out the mixture from locations in the chamber other than the bottom.

Steam pulsing - Air dilution by using a series of steam pulses, in which the chamber is alternately

pressurized and then depressurized to near atmospheric pressure.

Vacuum pumps - are to suck air or air/steam mixtures from the chamber.

Superatmospheric - This type of cycle uses a vacuum pump. It starts with a vacuum followed by a steam

pulse and then a vacuum followed by a steam pulse. The number of pulses depends on the particular

autoclave and cycle chosen.

Subatmospheric - Similar to superatmospheric cycles, but chamber pressure never exceeds atmospheric

until they pressurize up to the sterilizing temperature.

Autoclave quality assurance There are physical, chemical, and biological indicators that can be used to ensure an autoclave reaches

the correct temperature for the correct amount of time.

Chemical indicators can be found on medical packaging and autoclave tape, and these change

color once the correct conditions have been met. This color change indicates that the object inside the

package, or under the tape, has been processed. Biological indicators contain spores of a heat-resistant

bacterium, Geobacillus stearothermophilus. If the autoclave does not reach the right temperature, when

incubated the spores will germinate, and their metabolism will change the color of a pH-sensitive

chemical. Some physical indicators consist of an alloy designed to melt only after being subjected to the

relevant holding time. If the alloy melts, the change will be visible.

Some computer-controlled autoclaves use an F0 (F-nought) value to control the sterilization cycle.

F0 values are set as the number of minutes of equivalent sterilization at 121 °C (250 °F) at 15 psi

(100 kPa) above atmospheric pressure for 15 minutes. Since exact temperature control is difficult, the

temperature is monitored, and the sterilization time adjusted accordingly.

Standard operation procedure of autoclave

1. Check the water level and maintain it up to the mark /height of the stand 2. Ensure there is no leakage i.e. outlet is tightly closed 3. Place the articles with defined standards on through with indicator tape. Ensure date of

autoclave is mentioned over tape/ sticker

4. Close the lid and tighten the screw clamps 5. Ensure steam/ pressure release value must be open to release trapped air 6. Switch on the electric supply 7. Allow the steam to generate and release it for sometime to ensure that all air has been replaced

by steam 8. Close the steam release valve and allow the pressure to rise up to 15 lbs 9. Maintain 15 lbs pressure for 15-20 minutes 10. Switch off the electric supply 11. Allow the pressure to come down by its own while slowly releasing the valve 12. Open the lid of autoclave and take out the articles 13. Ensure indicator tape color is changed to brown color otherwise repeat the same procedure

Uses Autoclaves are widely used in microbiology, medicine, tattooing, body piercing, veterinary

science, mycology, dentistry, chiropody and prosthetic fabrication.

Typical loads include glassware, medical waste, utensils, animal cage bedding, and Lysogeny broth.

A notable growing application of autoclaves is in the treatment and sterilization of waste, such as

pathogenic hospital waste. Machines in this category largely operate under the same principles as the

original autoclave in that they are able to neutralize potentially infectious agents by utilizing pressurized

steam and superheated water.

2.4. Incubator

Incubator is an appliance shaped like a box which maintains the desired temperature inside it

Structure: (i) A box or container with insulated walls and a door fitted with a latch to close the door firmly. (ii) A hole in the center of its roof for insertion of a thermometer to read the temperature of the inside chamber. (iii) Its base contains a heating unit heated through electricity. (iv) In the front of the base on one side is a knob which can switch-on and switch-off the instrument. (v) On the backside is fitted a thermostat to regulate the desired temperature. (vi) In the centre of the front or besides the knob, a bulb is fitted to indicate whether the instrument is off or on (vii) The internal chamber is provided with one or more shelves.

Standard operation procedure of incubator

1. Wipe the racks of incubator with 75% alcohol before use 2. Switch on the electric supply 3. Set the optimal temperature by regulator for the growth of microbe 4. Place the articles on the rack 5. Petri dishes should be placed in inverted position 6. Place a 3/4th distilled water filled 100ml beaker in the incubator for maintaining the appropriate

moisture 7. Note down the time of placing the articles and let them incubate for required incubation period

8. Do not open the door in between the incubation period 9. On completion of incubation period remove the articles from the incubator 10. Switch off the incubator

Uses

1. To keep the microbial culture at desired temperature for growth 2. To keep section cutting material embedded in paraffin wax (at 50-55°C) 3. Incubate eggs: for this a dish containing water is kept inside the incubator to provide moisture

to the eggs. The eggs are rotated daily to prevent sticking of the embryo to the shell membranes. The temperature is maintained at about38°- 40°C.

4. To study the action of chemicals, enzymes etc. at different temperatures. 5. Specimens such as pinned and stretched insects can be dried in it, so that they are not spoiled.

2.5. UV – visible spectrophotometer

Molecules (e.g. DNA proteins etc.) absorb and emit electromagnetic radiation of particular wavelength. This property of molecules is used in spectrophotometry. Spectrophotometry is a technique which is widely used to measure the absorption of radiation in the visible and UV regions of the spectrum.

A spectrophotometer is employed to measure the amount of light that a sample absorbs. The instrument operates by passing a beam of light through a sample and measuring the intensity of light reaching a detector.

The beam of light consists of a stream of photons. When a photon encounters an analyte molecule (the analyte is the molecule being studied), there is a chance the analyte will absorb the photon. This absorption reduces the number of photons in the beam of light, thereby reducing the intensity of the light beam.

Structure of spectrophotometer

Experimental Procedure The following simulation illustrates the procedures for making spectrophotometric measurements.

• First, the intensity of light (I0) passing through a blank is measured. The intensity is the number of photons per second. The blank is a solution that is identical to the sample solution except that

the blank does not contain the solute that absorbs light. This measurement is necessary, because the cell itself scatters some of the light.

• Second, the intensity of light (I) passing through the sample solution is measured. (In practice, instruments measure the power rather than the intensity of the light. The power is the energy per second, which is the product of the intensity (photons per second) and the energy per photon.)

• Third, the experimental data is used to calculate two quantities: the transmittance (T) and the absorbance (A).

T = I

I0

A = - log10 T

The transmittance is simply the fraction of light in the original beam that passes through the sample and reaches the detector. The remainder of the light, 1 - T, is the fraction of the light absorbed by the sample. (Do not confuse the transmittance with the temperature, which often is given the symbol T.)

In most applications, one wishes to relate the amount of light absorbed to the concentration of the absorbing molecule. It turns out that the absorbance rather than the transmittance is most useful for this purpose. If no light is absorbed, the absorbance is zero (100% transmittance). Each unit in absorbance corresponds with an order of magnitude in the fraction of light transmitted. For A = 1, 10% of the light is transmitted (T = 0.10) and 90% is absorbed by the sample. For A = 2, 1% of the light is transmitted and 99% is absorbed. For A = 3, 0.1% of the light is transmitted and 99.9% is absorbed.

Using the simulation below, perform the following steps:

• Measure the intensity of light passing through the blank. • Measure the intensity of light passing through the sample. • Calculate the transmittance. • Calculate the absorbance.

Note: For each measurement, run the simulation long enough to detect at least 1000 photons. There is substantial random error in the intensity, and the more photons that are counted, the lower the relative uncertainty in the results.

Standard operation procedure of UV-visible spectrophotometer

1. Switch on the electric supply 2. Switch on the instrument 3. Allow the system to stabilize for 15-20 minutes 4. Switch on the required UV/visible lamp by toggle option on the screen 5. Calibrate the system using calibrate option on the screen 6. Select the desired mode i.e. single wavelength , multiple wavelength , range scan or time scan

according to experiment 7. Set the required parameters in desired mode 8. Place the cuvette with the reference solution in the slot provided with transparent side facing

the slit for light to pass, and take the reading of blank 9. Replace the reference cuvette with sample and click on the sample option on screen to take the

reading of the sample 10. For reassuring the reference in between the experiment the reference can be placed and taken

the reading using the reference option on the screen 11. On completion of work remove the cuvette from the instrument and clean it with distilled

water 12. Switch off the UV/ visible lamp 13. Switch off the program on the computer 14. Switch off the electric supply 15. Care should be taken not to switch on and off UV lamp frequently 16. Do not switch off system directly without switching off the lamp 17. Any spillage should be immediately cleaned 18. Use quartz cuvette only when working with UV lamp

Application of UV – visible spectrophotometer

Majorly two type of analysis are being done with UV – visible spectrophotometer

Quality analysis (through standard curve)

Quantity analysis (through spectrum)

Under the above two analyses following practical analysis are being done:

Chemical analysis

Food safety analysis

Blood analysis

DNA/RNA conc. analysis

Residual pesticides analysis

Residual chlorine analysis

2.6. Thermal cycler (PCR technique)

The thermal cycler (also known as a Thermocycler, PCR Machine or DNA Amplifier) is a

laboratory apparatus used to amplify segments of DNA via the polymerase chain reaction (PCR) process.

The device has a thermal block with holes where tubes holding the PCR reaction mixtures can be

inserted. The cycler then raises and lowers the temperature of the block in discrete, pre-programmed

steps.

The earliest thermal cyclers were designed for use with the Klenow fragment of DNA Polymerase I. Since

this enzyme is destroyed during each heating step of the amplification process, new enzyme had to be

added every cycle. This led to a cumbersome machine based on an automated pipettor, with open

reaction tubes. Later, the PCR process was adapted to the use of thermostable DNA

polymerase from Thermus aquaticus, which greatly simplified the design of the thermal cycler. While in

some old machines the block is submerged in oil bath to control temperature, in modern PCR machines

a Peltier element is commonly used. Quality thermal cyclers often contain silver blocks to achieve fast

temperature changes and uniform temperature throughout the block.

Modern thermal cyclers are equipped with a heated lid, a heated plate that presses against the lids of

the reaction tubes. This prevents condensation of water from the reaction mixtures on the insides of the

lids and makes it unnecessary to use PCR oil to cover the reaction mixture. Some thermal cyclers are

equipped with multiple blocks allowing several different PCR reactions to be carried out simultaneously.

Also some apparatus have a gradient function, which allows different temperatures in different parts of

the block. This is particularly useful when testing suitable annealing temperatures for primers.

Principle Developed in 1983 by Kary Mullis, PCR is now a common and often indispensable technique.

The polymerase chain reaction (PCR) is a technique in molecular biology to amplify a single or few copies

of a piece of DNA across several orders of magnitude, generating thousands to millions of copies of a

particular DNA sequence. The method relies on thermal cycling, consisting of cycles of repeated heating

and cooling of the reaction for DNA melting and enzymatic replication of the DNA. Primers (short DNA

fragments) containing sequences complementary to the target region along with a DNA

polymerase (after which the method is named) are key components to enable selective and repeated

amplification. As PCR progresses, the DNA generated is itself used as a template for replication, setting

in motion a chain reaction in which the DNA template is exponentially amplified. PCR can be extensively

modified to perform a wide array of genetic manipulations.

Almost all PCR applications employ a heat-stable DNA polymerase, such as Taq polymerase, an enzyme

originally isolated from the bacterium Thermus aquaticus. This DNA

polymerase enzymatically assembles a new DNA strand from DNA building blocks, the nucleotides, by

using single-stranded DNA as a template and DNA oligonucleotides (also called DNA primers), which are

required for initiation of DNA synthesis. The vast majority of PCR methods use thermal cycling, i.e.,

alternately heating and cooling the PCR sample to a defined series of temperature steps. These thermal

cycling steps are necessary first to physically separate the two strands in a DNA double helix at a high

temperature in a process called DNA melting. At a lower temperature, each strand is then used as

the template in DNA synthesis by the DNA polymerase to selectively amplify the target DNA. The

selectivity of PCR results from the use of primers that are complementary to the DNA region targeted

for amplification under specific thermal cycling conditions.

PCR 1.After DNA has been extracted from blood, cheek cells, hair, or anything else, put the DNA sample into a special PCR tube. 2. Add primer 1 to the tube. 3. Add primer 2 to the tube. 4. Add nucleotides solution to the tube. 5. Add DNA polymerase to the tube. 6. Put the tube in the DNA thermal cycler. (In this cycler, the temperature is raised and lowered at certain times in order to stimulate the specific steps. The temperature is raised, and this causes the DNA stands to separate. Then, it is cooled and the primers attach to the strands. The temperature is slightly raised again and this activates the DNA polymerase. Eventually, the target sequence separates and get copied several times. After 30 cycles, there are over 1 billion fragments of the target sequence.)

Standard operation procedure of thermal cycler

1. Switch on the electric supply 2. Switch on the thermal cycler 3. Press f1 key to display the program menu 4. Select the program by pressing f1 key 5. Select /enter the reaction volume 6. Unlock machine and open the lid 7. Place the sample in the space provided 8. Cover the lid and lock the machine 9. Select start by pressing f1 key 10. After completion of the selected program the temperature of machine would be automatically

set at 40C 11. Press stop 12. Again press stop to exit 13. Switch off the cycler 14. Switch off the electric supply 15. Unlock the machine and open the lid 16. Remove the sample 17. Clean the work area

Applications

1. DNA cloning for sequencing 2. DNA-based phylogeny 3. functional analysis of genes 4. the diagnosis of hereditary diseases 5. the identification of genetic fingerprints (used in forensic sciences and paternity testing) 6. the detection and diagnosis of infectious disease

2.7. pH meter

A pH meter is an electronic instrument used to measure the

special probes are sometimes used to measure the pH of semi

consists of a special measuring probe (a

measures and displays the pH reading. pH meter is nothing else but a precise voltameter connected to

the pH electrode - kind of ion selective electrode. Voltage produced by the pH electrode is proportional

to logarithm of the H+ activity (as described by the Nernst equation). pH meter voltameter display is

scaled in such a way that the displayed result of measurement is just the pH of the solution.

Commercial pH electrodes used in pH meters consist of a H+

just of very thin glass), and internal and external reference electrodes, usually combined in one housing.

Principle Every water solution contains H+ ions. Their concentration is one of the most important parameters

describing solution properties.

Concentrations of H+ can change in a very wide range; it can be 10

are inconvenient to use so to simplify things Danish biochemist Søren Sørensen developed in 1909

the pH scale and introduced pH definition

0.1

It is much easier to use pH definition and to say "pH of the solution is 4.1" than to use concentrations

as in "H+ concentration is 0.000079M".

pH meter

instrument used to measure the pH (acidity or alkalinity) of a

special probes are sometimes used to measure the pH of semi-solid substances). A typical pH meter

consists of a special measuring probe (a glass electrode) connected to an electronic meter that

measures and displays the pH reading. pH meter is nothing else but a precise voltameter connected to

kind of ion selective electrode. Voltage produced by the pH electrode is proportional

activity (as described by the Nernst equation). pH meter voltameter display is

scaled in such a way that the displayed result of measurement is just the pH of the solution.

Commercial pH electrodes used in pH meters consist of a H+ selective membrane (whic

just of very thin glass), and internal and external reference electrodes, usually combined in one housing.

ions. Their concentration is one of the most important parameters

can change in a very wide range; it can be 10 M as well as 10-15

are inconvenient to use so to simplify things Danish biochemist Søren Sørensen developed in 1909

pH definition - minus logarithm base 10 of [H+]:

It is much easier to use pH definition and to say "pH of the solution is 4.1" than to use concentrations

.000079M".

) of a liquid (though

solid substances). A typical pH meter

ected to an electronic meter that

measures and displays the pH reading. pH meter is nothing else but a precise voltameter connected to

kind of ion selective electrode. Voltage produced by the pH electrode is proportional

activity (as described by the Nernst equation). pH meter voltameter display is

scaled in such a way that the displayed result of measurement is just the pH of the solution.

e (which can be made

just of very thin glass), and internal and external reference electrodes, usually combined in one housing.

ions. Their concentration is one of the most important parameters

15 M. Such numbers

are inconvenient to use so to simplify things Danish biochemist Søren Sørensen developed in 1909

It is much easier to use pH definition and to say "pH of the solution is 4.1" than to use concentrations -

Not only H+ ions are present in every water solution. Also OH

concentration can change in the same very wide range. Thus it is also convenient to use similar

definition to describe [OH-]

0.2

Calibration Before use pH meter must be calibrated. Each pH electrode used for measurements is slightly different

and its characteristic changes with aging. Without proper pH meter calibration results will be usually off

by at least several tenths of the unit. pH meter ca

calibration buffers of exactly known pH (these can be bought as either solutions or in solid form) in

which pH electrode is dipped and pH meter indications are corrected. Depending on the pH meter type

it may either recognize buffer automatically and perform calibration procedure almost on its own (just

asking for buffer change when needed) or you will have to calibrate it using knobs

once each calibration step is completed. In both cases underlying principle is the same

parameters are set assuming linear dependence between pH and electrode voltage.

pH meter calibration is more often referred to a

device dependent, but electrode dependent.

The pH probe measures pH as the activity of

tip. The probe produces a small voltage (about 0.06 volt per pH unit) that is measured and displayed as

pH units by the meter.

The meter circuit is no more than a

The input impedance of the meter must be very high because of the high

to 1000 MΩ — of the glass electrode

meter usually consists of operational amplifiers

about -17. The inverting amplifier converts the small voltage produced by the probe (+0.059 volt/pH)

into pH units, which are then offset by seven volts to give a reading on the pH scale. For example:

At neutral pH (pH 7) the voltage at the probe's output is 0 volts. 0 * 17 + 7 = 7.

At basic pH, the voltage at the probe's output ranges from +0 to +0.41 volts (7 * 0.

sample of pH 10 (3 pH units above neutral), 3 * 0.059 = 0.18 volts), the output of the meter's amplifier is

0.18 * 17 + 7 = 10.

At acid pH, the voltage at the probe's output ranges from

units below neutral), -3 * 0.059 = -0.18 volts, the output of the meter's amplifier is

The two basic adjustments performed at calibration set the gain and offset of the inverting amplifier.

ions are present in every water solution. Also OH- ions are always present, and their

concentration can change in the same very wide range. Thus it is also convenient to use similar

re use pH meter must be calibrated. Each pH electrode used for measurements is slightly different

and its characteristic changes with aging. Without proper pH meter calibration results will be usually off

by at least several tenths of the unit. pH meter calibration procedure calls for use of two or three

of exactly known pH (these can be bought as either solutions or in solid form) in

and pH meter indications are corrected. Depending on the pH meter type

it may either recognize buffer automatically and perform calibration procedure almost on its own (just

asking for buffer change when needed) or you will have to calibrate it using knobs and changing buffers

once each calibration step is completed. In both cases underlying principle is the same

parameters are set assuming linear dependence between pH and electrode voltage.

pH meter calibration is more often referred to as pH electrode calibration, as parameters set are not

device dependent, but electrode dependent.

The pH probe measures pH as the activity of hydrogen ions surrounding a thin-walled glass bulb at its

tip. The probe produces a small voltage (about 0.06 volt per pH unit) that is measured and displayed as

The meter circuit is no more than a voltmeter that displays measurements in pH units instead of volts.

must be very high because of the high resistance —

glass electrode probes typically used with pH meters. The circuit of a simple pH

operational amplifiers in an inverting configuration, with a total voltage

17. The inverting amplifier converts the small voltage produced by the probe (+0.059 volt/pH)

ts, which are then offset by seven volts to give a reading on the pH scale. For example:

At neutral pH (pH 7) the voltage at the probe's output is 0 volts. 0 * 17 + 7 = 7.

At basic pH, the voltage at the probe's output ranges from +0 to +0.41 volts (7 * 0.059 = 0.41). So for a

sample of pH 10 (3 pH units above neutral), 3 * 0.059 = 0.18 volts), the output of the meter's amplifier is

At acid pH, the voltage at the probe's output ranges from -0.41 volts to -0. So for a sample of pH 4 (3 pH

0.18 volts, the output of the meter's amplifier is -0.18 * 17 + 7 = 4.

The two basic adjustments performed at calibration set the gain and offset of the inverting amplifier.

ions are always present, and their

concentration can change in the same very wide range. Thus it is also convenient to use similar

re use pH meter must be calibrated. Each pH electrode used for measurements is slightly different

and its characteristic changes with aging. Without proper pH meter calibration results will be usually off

libration procedure calls for use of two or three pH

of exactly known pH (these can be bought as either solutions or in solid form) in

and pH meter indications are corrected. Depending on the pH meter type

it may either recognize buffer automatically and perform calibration procedure almost on its own (just

and changing buffers

once each calibration step is completed. In both cases underlying principle is the same - gain and offset

, as parameters set are not

walled glass bulb at its

tip. The probe produces a small voltage (about 0.06 volt per pH unit) that is measured and displayed as

that displays measurements in pH units instead of volts.

— approximately 20

probes typically used with pH meters. The circuit of a simple pH

in an inverting configuration, with a total voltage gain of

17. The inverting amplifier converts the small voltage produced by the probe (+0.059 volt/pH)

ts, which are then offset by seven volts to give a reading on the pH scale. For example:

059 = 0.41). So for a

sample of pH 10 (3 pH units above neutral), 3 * 0.059 = 0.18 volts), the output of the meter's amplifier is

0. So for a sample of pH 4 (3 pH

0.18 * 17 + 7 = 4.

The two basic adjustments performed at calibration set the gain and offset of the inverting amplifier.

History The first commercial pH meters were built around 1936 by Radiometer in Denmark and by Arnold Orville

Beckman in the United States. While Beckman was an assistant professor of chemistry at the California

Institute of Technology, he was asked to devise a quick and accurate method for measuring the acidity

of lemon juice for the California Fruit Growers Exchange (Sunkist). Beckman's invention helped him to

launch the Beckman Instruments company (now Beckman Coulter). In 2004 the Beckman pH meter was

designated an ACS National Historical Chemical Landmark in recognition of its significance as the first

commercially successful electronic pH meter.[2]

In the 1970s Jenco Electronics of Taiwan designed and manufactured the first portable digital pH meter.

This meter was sold under Cole-Palmer's label.

Standard operation procedure of pH meter

1. Check that electrode is well kept in distilled water 2. Switch on the electric supply 3. Switch on pH meter 4. Allow the system to stabilize for the 15 min. prior to use 5. Calibrate the equipment with the standard buffer solution of pH 4, followed by pH 7 and then

pH 9.2. After every solution, electrode has to be washed by distilled water. Buffer solution should be at room temperature

6. Check the pH of your sample 7. Switch off the pH meter 8. Wash and wipe the electrode 9. Switch off the electric supply 10. Clean the work area

2.8. Vortex

A vortex (plural: vortices) is a spinning, often turbulent, flow of fluid. Any spiral motion with closed

streamlines is vortex flow. The motion of the fluid swirling rapidly around a center is called a vortex. The

speed and rate of rotation of the fluid in a free (irrotational) vortex are greatest at the center, and

decrease progressively with distance from the center, whereas the speed of a forced (rotational) vortex

is zero at the center and increases proportional to the distance from the center. Both types of vortices

exhibit a pressure minimum at the center, though the pressure minimum in a free vortex is much lower.

Vortices display some special properties:

The fluid pressure in a vortex is lowest in the center and rises progressively with distance from the

center. This is in accordance with Bernoulli's Principle. The core of a vortex in air is sometimes visible

because of a plume of water vapor caused by condensation in the low pressure of the core. The spout of

a tornado is a classic and frightening example of the visible core of a vortex. A dust devil is also the core

of a vortex, made visible by the dust drawn upwards by the turbulent flow of air from ground level into

the low pressure core.

The core of every vortex can be considered to contain a vortex line, and every particle in the vortex can

be considered to be circulating around the vortex line. Vortex lines can start and end at the boundary of

the fluid or form closed loops. They cannot start or end in the fluid. (See Helmholtz's theorems.) Vortices

readily deflect and attach themselves to a solid surface. For example, a vortex usually forms ahead of

the propeller disk or jet engine of a slow-moving airplane. One end of the vortex line is attached to the

propeller disk or jet engine, but when the airplane is taxiing the other end of the vortex line readily

attaches itself to the ground rather than end in midair. The vortex can suck water and small stones into

the core and then into the propeller disk or jet engine.

Two or more vortices that are approximately parallel and circulating in the same direction will merge to

form a single vortex. The circulation of the merged vortex will equal the sum of the circulations of the

constituent vortices. For example, a sheet of small vortices flows from the trailing edge of the wing or

propeller of an airplane when the wing is developing lift or the propeller is developing thrust. In less

than one wing chord downstream of the trailing edge of the wing these small vortices merge to form a

single vortex. If viewed from the tail of the airplane, looking forward in the direction of flight, there is

one wingtip vortex trailing from the left-hand wing and circulating clockwise, and another wingtip vortex

trailing from the right-hand wing and circulating anti-clockwise. The result is a region of downwash

behind the wing, between the pair of wingtip vortices. These two wingtip vortices do not merge because

they are circulating in opposite directions.

Vortices contain a lot of energy in the circular motion of the fluid. In an ideal fluid this energy can never

be dissipated and the vortex would persist forever. However, real fluids exhibit viscosity and this

dissipates energy very slowly from the core of the vortex. It is only through dissipation of a vortex due to

viscosity that a vortex line can end in the fluid, rather than at the boundary of the fluid. For example, the

wingtip vortices from an airplane dissipate slowly and linger in the atmosphere long after the airplane

has passed. This is a hazard to other aircraft and is known as wake turbulence.

Standard operation procedure of vortex

1. Switch on the electric supply 2. Place the sample /article vials in the slots provided on vortex 3. Vortex the vials by switching on the “ constant on”/ “touch on” system 4. Regulate the speed by “speed control” button 5. After completion , switch off “constant on” button 6. Switch off the electric supply

2.9. Dry bath

The versatile dry baths provide controlled dry heat for a wide variety of clinical and general chemistry

applications. The newly designed unparalleled accuracy, ratability, and flexibility in application. It has

unique auto tuned microprocessor based temperature controller witch controls the key parameters of

temperature. While granting user more control, than any other block heater for the rate [cost]. It also

has fully stainless steel body. Blocks are of aluminum anodized and autoclavable.

Standard operation procedure of dry bath

1. Switch on the electric supply 2. Switch on the dry bath 3. Place the “temperature controlled electrode” in the space provided on dry bath 4. Set the desired temperature by “set temp” button 5. Temperature may be increased or decreased by pressing arrow keys 6. Place the sample/article vials in the slots of dry bath 7. Note the time of placing the sample 8. After completion of the required incubation period; switch off the dry bath 9. Remove the vial from the system 10. Switch off the electric supply

Applications Dry bath holds very importance in following events:

FISH (Fluorescent in Situ Hybridization)

Southern, Northern, Western Blots

Microarray Hybridization

E-coli testing in Food, water- wastewater

ASTM C151 Cement testing

Cell Culture (Denaturing or defrosting cell lines)

COD Tests- wastewater

Histology- Hematology, Slide preparation

Viscosity Testing, ASTM D 4212

Microbiology applications

Bacteriology applications

PCR, Samples prep

Blood, Hematology, Sample prep

2.10. Hot plate

A hot plate is a self-contained tabletop small appliance that features single or double electric heating

elements and is thus portable. A hot plate is intended as a substitute for one of the burners from

an oven range or the cook top of a stove. Hot plates are often used in food preparation, generally for

small dishes in locations where a full kitchen stove would not be convenient.

In laboratory settings, hot plates are generally used to heat glassware or its contents. Some hotplates

also contain a magnetic stirrer, allowing the heated liquid to be stirred automatically.

Standard operation procedure of hot plate

1. Switch on the electric supply 2. Allow the machine to warm up 3. Place the gauge on the heater 4. Place your article on the gauge for the required time 5. Periodical shaking is required to avoid bubbling and busting 6. Switch off the electric supply 7. Remove the article 8. Clean the work area 9. Avoid bubbling and spillage

2.11. Digital weighing machine

An analytical balance is an instrument that is used to measure mass to a very high degree of precision. The weighing pan of a high precision(0.01mg) are inside a transparent enclosure with doors so dust does not collect and so any currents in the room do not affect the delicate balance. The sample must be at room temperature to prevent natural convection from forming air currents.

Standard operation procedure of digital weighing machine

1. Switch on the electric supply 2. Switch on balance 3. Allow the system to stabilize 4. Place the butter paper on balance 5. Set the machine in “Re-zero” position by pressing “tare” 6. Weigh the chemicals in required amount 7. Switch off balance and then electric supply 8. Clean the weighing bowl and balance cabinet 9. Fan should be switched off while weighing the reagents

2.12. Electrophoresis unit

Horizontal and Vertical Gel Systems

A gel electrophoresis apparatus must allow the researcher to maintain a uniform electric field across the gel, provide cooling to prevent thermal artifacts, and allow access to the gel for sample loading and monitoring the run. Two types of apparatus are in common use: vertical and horizontal. Vertical gel systems are further subdivided into slab gels and tube gels. In general, agarose gels are run in the horizontal format, while acrylamide gels are run vertically.

The Horizontal Gel System

In its simplest form, a horizontal gel apparatus consists of a box which is divided into two compartments by a platform in the middle. The gel is placed on this platform, and buffer is added until the gel is fully submerged. Electrodes in each compartment supply the electric field. The resulting current flows through both the gel and the buffer over the gel, so the thickness of these must be controlled for fully reproducible results. Cooling is provided by the buffer which surrounds the gel, and this buffer is often recirculated to prevent the development of a pH gradient and also to aid in temperature control. Access to the gel is through the overlaying buffer. Samples are loaded through this buffer layer, and the apparatus has a clear lid to allow the run to be monitored.

The Horizontal Gel Electrophoresis Apparatus. The gel rests on a platform which divides the apparatus into two chambers. Note that the buffer level is higher than the surface of the gel, so the two buffer chambers are connected. Samples loaded into the wells will migrate toward

the positive electrode.

One major limitation of the horizontal apparatus is that, since the two compartments are connected by a layer of buffer, it is not possible to use discontinuous buffer systems. Another limitation is that the gels are cast in trays which are not covered. Because atmospheric oxygen has full access to the upper surface of the gel, acrylamide will not polymerize in this system. Horizontal systems are primarily used to run agarose gels which are run in continuous buffer systems and are not affected by O2. For agarose gels, the

simplicity and ease of use of the horizontal system make this system the best choice.

Casting a horizontal agarose gel

Horizontal gels are cast in trays which have removable ends. The ends are installed (often the ends are simply adhesive tape), and the molten agarose solution is poured into the tray. A comb is inserted so that the teeth penetrate the gel to within 1-2mm of the bottom of the tray. The overall gel thickness is generally 0.5-1cm. The gel is allowed to cool, the comb is removed, and the gel is mounted in the apparatus

Vertical Gel Systems

Slab gels

A typical vertical apparatus used for sequencing is shown in the figure below. This system shows the components common to all vertical slab systems. The gel is cast between two glass plates, separated by spacers, typically <2mm thick. The gel is mounted in the system so that the top is in contact with the negative electrode chamber, and the bottom is in contact with the positive electrode chamber. Unlike the horizontal system, the only connection between the buffer chambers is through the gel. This allows precise and reproducible control of the voltage gradient. Because of the high resistance of the thin gel, the apparatus must have provisions for cooling. In the system shown, the front of the gel cassette is exposed to the air, while the back of the gel is held against a metal plate which dissipates heat rapidly. In some systems, the upper buffer chamber extends almost to the bottom of the gel, and the upper buffer is used for cooling. The relatively small amount of current carried through the gel means that buffer recirculation is generally not required.

The Vertical Gel Electrophoresis Apparatus.

The gel is clamped into the apparatus so that the lower end is immersed in the lower buffer chamber, and the upper end forms one wall of the upper chamber. The gel provides the only electrical connection between the two buffer chambers. Cooling is provided by a metal heat sink behind the gel. This type of apparatus allows high voltages (>1500V) to be used. The Vertical Gel Electrophoresis Apparatus. The gel is clamped into the apparatus so that the lower end is immersed in the lower buffer chamber, and the upper end forms one wall of the upper chamber. The gel provides the only electrical connection betweenthe two buffer chambers. Cooling is provided by a metal heat sink behind the gel. This type of apparatus allows high voltages (>1500V) to be used.

The figure below shows a standard "mini" gel apparatus. Such gels, generally 10cm x 10cm or smaller, have become the standard for many applications, because of their ease of preparation and handling, and short run times. As with the "full size" system, the gels are cast between glass plates, but in the mini-gel system, the cassettes are mounted onto a "U" shaped frame, so that the cassettes themselves form the sides of the negative electrode chamber. This assembly is placed in a tank of buffer which contains the positive electrode. This means that the gels are effectively submerged in buffer during the run, providing optimal cooling.

The vertical mini-gel apparatus.

The upper (negative) electrode chamber is formed by the two gel cassettes, clamped to the "core". This assembly is submerged in the lower (positive) electrode buffer chamber, which provides cooling. As with the full size vertical system, the gels provide the only electrical connection between the two chambers.

In general, vertical slab gels are loaded through the top, under a layer of buffer. The gels are monitored during the run through the front glass plate. The fact that the body of the gel in these systems cannot be accessed until the end of the run can be an inconvenience. Some sample recovery techniques used on horizontal gels are not available for vertical gels. However, the resolution and reproducibility of vertical polyacrylamide gels more than compensate for this. Casting a vertical slab gel

Vertical gels are cast in a cassette made up of two glass plates separated by spacers who run along the sides of the plates. The bottom of the cassette is sealed by some temporary means (tape, agarose, or a gasket). The gel monomer solution is treated to initiate polymerization, and poured into the cassette. A comb is inserted into the top of the cassette to form the sample wells, and the gel is allowed sufficient time to polymerize. After polymerization, the bottom of the gel is unsealed, and the cassette is mounted in the apparatus.

Principle:

Electrophoresis is a technique used to separate and sometimes purify macromolecules - especially proteins and nucleic acids - that differ in size, charge or conformation. As such, it is one of the most widely-used techniques in biochemistry and molecular biology.

When charged molecules are placed in an electric field, they migrate toward either the positive or negative pole according to their charge. In contrast to proteins, which can have either a net positive or net negative charge, nucleic acids have a consistent negative charge imparted by their phosphate backbone, and migrate toward the anode.

Proteins and nucleic acids are electrophoresed within a matrix or "gel". Most commonly, the gel is cast in the shape of a thin slab, with wells for loading the sample. The gel is immersed within an electrophoresis buffer that provides ions to carry a current and some type of buffer to maintain the pH at a relatively constant value.

The gel itself is composed of either agarose or polyacrylamide, each of which has attributes suitable to particular tasks:

Agarose is a polysaccharide extracted from seaweed. It is typically used at concentrations of 0.5 to 2%. The higher the agarose concentration the "stiffer" the gel. Agarose gels are extremely easy to prepare: you simply mix agarose powder with buffer solution, melt it by heating, and pour the gel. It is also non-toxic.

Agarose gels have a large range of separation, but relatively low resolving power. By varying the concentration of agarose, fragments of DNA from about 200 to 50,000 bp can be separated using standard electrophoretic techniques.

Polyacrylamide is a cross-linked polymer of acrylamide. The length of the polymer chains is dictated by the concentration of acrylamide used, which is typically between 3.5 and 20%. Polyacrylamide gels are significantly more annoying to prepare than agarose gels. Because oxygen inhibits the polymerization process, they must be poured between glass plates (or cylinders).

Acrylamide is a potent neurotoxin and should be handled with care! Wear disposable gloves when handling solutions of acrylamide, and a mask when weighing out powder. Polyacrylamide is considered to be non-toxic, but polyacrylamide gels should also be handled with gloves due to the possible presence of free acrylamide.

Polyacrylamide gels have a rather small range of separation, but very high resolving power. In the case of DNA, polyacrylamide is used for separating fragments of less than about 500 bp. However, under appropriate conditions, fragments of DNA differing is length by a single base pair are easily resolved. In contrast to agarose, polyacrylamide gels are used extensively for separating and characterizing mixtures of proteins.

Nucleic acids

An agarose gel of a PCR product compared to a DNA ladder.

In the case of nucleic acids, the direction of migration, from negative to positive electrodes, is due to the

naturally-occurring negative charge carried by their sugar-phosphate backbone.

Double-stranded DNA fragments naturally behave as long rods, so their migration through the gel is

relative to their size or, for cyclic fragments, their radius of gyration. Single-stranded DNA or RNA tends

to fold up into molecules with complex shapes and migrate through the gel in a complicated manner

based on their tertiary structure. Therefore, agents that disrupt the hydrogen bonds, such as sodium

hydroxide or formamide, are used to denature the nucleic acids and cause them to behave as long rods

again.

Gel electrophoresis of large DNA or RNA is usually done by agarose gel electrophoresis. See the "Chain

termination method" page for an example of a polyacrylamide DNA sequencing gel. Characterization

through ligand interaction of nucleic acids or fragments may be performed by mobility shift affinity

electrophoresis.

Electrophoresis of RNA samples can be used to check for genomic DNA contamination and also for RNA

degradation. RNA from eukaryotic organisms shows distinct bands of 28s and 18s rRNA, the 28s band

being approximately twice as intense as the 18s band. Degraded RNA has less sharply defined bands, has

a smeared appearance, and intensity ratio is less than 2:1.

Proteins

SDS-PAGE autoradiography - The indicated proteins are present in different concentrations in the two samples.

Proteins, unlike nucleic acids, can have varying charges and complex shapes; therefore they may not

migrate into the polyacrylamide gel at similar rates, or at all, when placing a negative to positive EMF on

the sample. Proteins therefore, are usually denatured in the presence of a detergent such as sodium

Dodecyl sulfate/sodium Dodecyl phosphate (SDS/SDP) that coats the proteins with a negative

charge. Generally, the amount of SDS bound is relative to the size of the protein (usually 1.4g SDS per

gram of protein), so that the resulting denatured proteins have an overall negative charge, and all the

proteins have a similar charge to mass ratio. Since denatured proteins act like long rods instead of

having a complex tertiary shape, the rate at which the resulting SDS coated proteins migrate in the gel is

relative only to its size and not its charge or shape.

Proteins are usually analyzed by sodium Dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE),

by native gel electrophoresis, by quantitative preparative native continuous polyacrylamide gel

electrophoresis (QPNC-PAGE), or by 2-D electrophoresis.

Characterization through ligand interaction may be performed by electroblotting or by affinity

electrophoresis in agarose or by capillary electrophoresis as for estimation of binding constants and

determination of structural features like glycan content through lectin binding.

Preparing and Running Standard Agarose DNA Gels

The equipment and supplies necessary for conducting agarose gel electrophoresis are relatively simple

and include:

1. An electrophoresis chamber and power supply

2. Gel casting trays, which are available in a variety of sizes and composed of UV-

transparent plastic. The open ends of the trays are closed with tape while the gel is

being cast, then removed prior to electrophoresis.

3. Sample combs, around which molten agarose is poured to form sample wells in the gel.

4. Electrophoresis buffer, usually Tris-acetate-EDTA (TAE) or Tris-borate-EDTA (TBE).

5. Loading buffer, which contains something dense (e.g. glycerol) to allow the sample to

"fall" into the sample wells, and one or two tracking dyes, which migrate in the gel and

allow visual monitoring or how far the electrophoresis has proceeded.

6. Ethidium bromide, a fluorescent dye used for staining nucleic acids. NOTE: Ethidium

bromide is a known mutagen and should be handled as a hazardous chemical - wear

gloves while handling.

7. Transilluminator (an ultraviolet light box), which is used to visualize Ethidium bromide-

stained DNA in gels. NOTE: always wear protective eyewear when observing DNA on a

Transilluminator to prevent damage to the eyes from UV light.

To pour a gel, agarose powder is mixed with electrophoresis buffer to the desired concentration, and then heated in a microwave oven until completely melted. Most commonly, Ethidium bromide is added to the gel (final concentration 0.5 ug/ml) at this point to facilitate visualization of DNA after electrophoresis. After cooling the solution to about 60C, it is poured into a casting tray containing a sample comb and allowed to solidify at room temperature or, if you are in a big hurry, in a refrigerator.

After the gel has solidified, the comb is removed, using care not to rip the bottom of the wells. The gel, still in its plastic tray, is inserted horizontally into the electrophoresis chamber and just covered with buffer. Samples containing DNA mixed with loading buffer are then pipeted into the sample wells, the lid and power leads are placed on the apparatus, and a current is applied. You can confirm that current is flowing by observing bubbles coming off the electrodes.DNA will migrate towards the positive electrode, which is usually colored red.

The distance DNA has migrated in the gel can be judged by visually monitoring migration of the tracking dyes. Bromophenol blue and xylene cyanol dyes migrate through agarose gels at roughly the same rate as double-stranded DNA fragments of 300 and 4000 bp, respectively.

When adequate migration has occurred, DNA fragments are visualized by staining with Ethidium bromide. This fluorescent dye intercalates between bases of DNA and RNA. It is often incorporated into the gel so that staining occurs during electrophoresis, but the gel can also be stained after electrophoresis by soaking in a dilute solution of Ethidium bromide. To visualize DNA or RNA, the gel is placed on an ultraviolet Transilluminator. Be aware that DNA will diffuse within the gel over time, and examination or photography should take place shortly after cessation of electrophoresis.

Migration of DNA Fragments in Agarose

Fragments of linear DNA migrate through agarose gels with a mobility that is inversely proportional to

the log10 of their molecular weight. In other words, if you plot the distance from the well that DNA

fragments have migrated against the log10 of either their molecular weights or number of base pairs, a

roughly straight line will appear.

Circular forms of DNA migrate in agarose distinctly differently from linear DNAs of the same mass. Typically, uncut plasmids will appear to migrate more rapidly than the same plasmid when linearized. Additionally, most preparations of uncut plasmid contain at least two topologically-different forms of DNA, corresponding to super coiled forms and nicked circles. The image to the right shows an Ethidium-stained gel with uncut plasmid in the left lane and the same plasmid linearized at a single site in the right lane.

Several additional factors have important effects on the mobility of DNA fragments in agarose gels, and can be used to your advantage in optimizing separation of DNA fragments. Chief among these factors are:

Agarose Concentration: By using gels with different concentrations of agarose, one can resolve different sizes of DNA fragments. Higher concentrations of agarose facilite separation of small DNAs, while low agarose concentrations allow resolution of larger DNAs.

The image to the right shows migration of a set of DNA fragments in three concentrations of agarose, all of which were in the same gel tray and electrophoresed at the same voltage and for identical times. Notice how the larger fragments are much better resolved in the 0.7% gel, while the small fragments separated best in 1.5% agarose. The 1000 bp fragment is indicated in each lane.

Voltage: As the voltage applied to a gel is increased, larger fragments migrate proportionally faster that small fragment. For that reason, the best resolution of fragments larger than about 2 kb is attained by applying no more than 5 volts per cm to the gel (the cm value is the distance between the two electrodes, not the length of the gel).

Electrophoresis Buffer: Several different buffers have been recommended for electrophoresis of DNA. The most commonly used for duplex DNA are TAE (Tris-acetate-EDTA) and TBE (Tris-borate-EDTA). DNA fragments will migrate at somewhat different rates in these two buffers due to differences in ionic strength. Buffers not only establish a pH, but provide ions to support conductivity. If you mistakenly use water instead of buffer, there will be essentially no migration of DNA in the gel! Conversely, if you use concentrated buffer (e.g. a 10X stock solution), enough heat may be generated in the gel to melt it.

Effects of Ethidium Bromide: Ethidium bromide is a fluorescent dye that intercalates between bases of nucleic acids and allows very convenient detection of DNA fragments in gels, as shown by all the images on this page. As described above, it can be incorporated into agarose gels, or added to samples of DNA before loading to enable visualization of the fragments within the gel. As might be expected, binding of Ethidium bromide to DNA alters its mass and rigidity, and therefore its mobility.

Other Considerations

Agarose gels, as discussed above provide the most commonly-used means of isolating and purifying fragments of DNA, which is a prerequisite for building any type of recombinant DNA molecule.

By varying buffer composition and running conditions, the utility of agarose gels can be extended. Examples include:

• Pulsed field electrophoresis is a technique in which the direction of current flow in the electrophoresis chamber is periodically altered. This allows fractionation of pieces of DNA ranging from 50,000 to 5 million bp, which is much larger than can be resolved on standard gels.

• Alkaline agarose gels are prepared with and electrophoresed in buffers containing sodium hydroxide. Such alkaline conditions are useful for analyzing single-stranded DNA.

History

§ 1930s - first reports of the use of sucrose for gel electrophoresis

§ 1955 - introduction of starch gels, mediocre separation

§ 1959 - introduction of acrylamide gels; disc electrophoresis (Ornstein and Davis); accurate control of

parameters such as pore size and stability; and (Raymond and Weintraub)

§ 1969 - introduction of denaturing agents especially SDS separation of protein subunit (Weber and

Osborn)

§ 1970 - Laemmli separated 28 components of T4 phage using a stacking gel and SDS

§ 1975 - 2-dimensional gels (O’Farrell); isoelectric focusing then SDS gel electrophoresis

§ 1977 - sequencing gels

§ late 1970s - agarose gels

§ 1983 - pulsed field gel electrophoresis enables separation of large DNA molecules

§ 1983 - introduction of capillary electrophoresis

A 1959 book on electrophoresis by Milan Bier cites references from the 1800s. However, Oliver

Smithies made significant contributions. Bier states: "The method of Smithies ... is finding wide application

because of its unique separatory power." Taken in context, Bier clearly implies that Smithies' method is

an improvement.

Standard operation procedure of vertical electrophoresis unit

1. Wipe the electrophoresis chamber with 75% alcohol 2. Place the casted gel slab along with the glass plate in the central part of electrophoresis tank 3. Pour the buffer solution in the gel tank till so that the upper and lower buffer tank are filled

completely 4. Cover the unit for the chamber saturation 5. Connect the power cord to the electrophoresis power supply according to the convention i.e.

red on anode and black on cathode 6. Load the sample along with control in the well of gel slab in the desired order with the help of

micropipette 7. Set the voltage to desired volt and switch on the power supply 8. Switch off the power of electrophoresis power supply when the sample from the wells reaches

3/4th of the gel 9. Carefully remove the gel slab from gel tank and use for next operation 10. Recollect the buffer from gel tank and store in freeze , it can be reused 11. Wipe the electrophoresis chamber with 75% alcohol 12. Care should be taken to remove the sealing agent used for casting gel from the glass slab before

starting the run

Standard operation procedure of horizontal electrophoresis unit

1. Wipe the electrophoresis chamber with 75% alcohol 2. Place the casted gel slab along with the glass plate in the central part of electrophoresis tank 3. Pour the buffer solution in the gel tank till the buffer level stands at 0.5 cm to 0.8cm above the

gel surface 4. Cover the unit for the chamber saturation 5. Connect the power cord to the electrophoresis power supply according to the convention i.e.

red on anode and black on cathode 6. Load the sample along with control in the well of gel slab in the desired order with the help of

micropipette 7. Set the voltage to desired volt and switch on the power supply 8. Switch off the power of electrophoresis power supply when the sample from the wells reaches

3/4th of the gel 9. Carefully remove the gel slab from gel tank and use for next operation 10. Recollect the buffer from gel tank and store in freeze , it can be reused 11. Wipe the electrophoresis chamber with 75% alcohol

2.13. UV Transilluminator

UV Transilluminators are designed for viewing DNA/RNA in agarose gels stained with Ethidium bromide (EB) or other dyes. Transparent hinged cover, with UV-blocking material to protect users from exposure to hazardous UV radiation, is mounted by slip-hinges for easy removal and replacement. These 302nm or 365nm UV lamps produce even distribution at the filter surface for better identification and documentation of viewed gels. The 302nm UV induces high fluorescence and sensitivity in Ethidium bromide/DNA complexes producing sharp, distinct bands. At this wavelength, photo-nicking, dimerization and photo-bleaching to gels are minimized. Unsurpassed UV intensity and outstanding irradiance uniformity enable visualization of even trace amounts of DNA. Airproof frame reduces the chance of gel damage or contamination.

The UV Transilluminator can be widely used in the laboratory of university, research department and corporation involved in molecular biology, molecular genetics, medical and sanitation, agriculture and other life science fields.

Standard operation procedure of UV Transilluminator

1. Switch on the electric supply 2. Switch on the Transilluminator 3. Place the gel slab on Transilluminator platform 4. Wear the provided UV shield

5. Switch on the UV lamp 6. Interpret the result 7. Switch off the UV lamp 8. Clean the platform with 75% alcohol 9. Switch off the electric supply

Exercise - 1

Preparation of buffer and reagents

Important definitions:

Reagent: complex mixture of chemicals generally used for chemical analysis

Solution: a combination of solute and solvent in a fixed proportion is called solution

Solute: dissolved substance, less in quantity

Solvent: dissolving substance, more in quantity

Stock solution: these are the solutions having a concentration of many folds higher than that actually required in the experiment

Working solution: these are the solutions prepared freshly from stock solution by diluting it as per the desired concentration

Standard solution: these are the solutions of known concentration and used for qualitative determination of a particular substance

Saturated solution: in these solution the amount of solute is in excess of that which can be completely dissolved at a given temperature is in equilibrium with the excess of undissolved solute

Buffer solution: A buffer solution is an aqueous solution consisting of a mixture of a weak acid and its conjugate base or a weak base and its conjugate acid. It has the property that the pH of the solution changes very little when a small amount of strong acid or base is added to it. Buffer solutions are used as a means of keeping pH at a nearly constant value in a wide variety of chemical applications

Percent solution: There are three types of percent solutions. All are parts of solute per 100 total parts of solution

Based on the following definitions you may calculate the concentration of a solution or calculate how to make up a specific concentration

1. % W/W -Percent of weight of solute in the total weight of the solution. Percent here is the number of grams of solute in 100 grams of solution

Example: A 100% (W/W) NaCl solution is made by weighing 100 g NaCl and dissolving in 100 g of solution

2. % W/V -Percent of weight of solution in the total volume of solution. Percent here is the number of grams of solute in 100mL of solution. This is probably the least significant way of naming a solution, but the most common way of doing it. In fact, any percent solution not stipulated as W/W, W/V, or V/V is assumed to be % W/V

Example: A 4% (W/V) NaCl solution is 4 g of NaCl in 100mL of solution

3. % V/V -Percent of volume of solute in the total volume of solution %V/V. Percent here is the number of milliliters of solute in 100 mL of solution

Example: A 10% (V/V) ethanol solution is 10 mL of ethanol in 100 mL of solution; unless otherwise stated, water is the solvent

MOLAR SOLUTIONS (M)

The definition of molar solution is a solution that contains 1 mole of solute in each liter of solution. A mole is the number of gram molecular weights (gmw). Therefore, we can also say a 1M = 1 GMW solute/liter solution.

1M NaCl solution would be

Na = MW of 23

Cl = MW of 35.5

NaCl = MW of 58.5

1M = 58.5 g of NaCl in 1 liter of solution

NORMAL SOLUTIONS:

The definition of a normal solution is a solution that contains 1 gram equivalent weight (gEW) per liter solution. An equivalent weight is equal to the molecular weight divided by the valence (replaceable H ions).

1N NaCl = 58.5 g/L

1N HCl = 36.5 g/L

1N H2S04 = 49 g/L

Problems involving normality are worked the same as those involving molarity but the valence must be considered:

1N HCL the MW= 36.5 the EW = 36.5 and 1N would be 36.5 g/L

1N H2SO4 the MW = 98 the EW = 49 and 1N would be 49 g/L

1N H3PO4 the MW = 98 the EW = 32.7 and 1N would be 32.7 g/L

MOLAL SOLUTIONS: A solution which contains one mole of the solute per 1000 grams of the solvent. For instance: adding 1.0 mole of solute to 2.0 kilograms of solvent constitutes a solution with a molality of 0.50 mol/kg. Such a solution may be described as "0.50 molal"

Preparation of buffer and reagents

a. 1M sodium hydroxide

Resuspend 40 gm of NaOH in 1000ml of distilled water and dissolve well. The solution is to be autoclaved at 121oC and 15lbs for 20 mins.

This solution can be kept for 15-20 days.

b. 0.1M sodium citrate

Resuspend 29.4gm of sodium citrate in 1000ml of distilled water and dissolve well. The solution is to be autoclaved at 121oC and 15lbs for 20 mins.

c. Tris EDTA

Tris 1M (pH 7.5)

Resuspend 121.4gm of Tris in 1000ml of distilled water and dissolve well.

EDTA 0.5M

Resuspend 186.4gm of EDTA in 1000ml of distilled water and dissolve well. EDTA requires constant stirring and mild heating and will dissolve completely when pH of the solution is 8

Working TE buffer

Final volume = 1000ml

Tris 10mM – 10ml from 1M stock

EDTA 1mM – 2ml from 0.5M stock

d. TNE buffer (10x) pH- 7.4 , 800ml

100mM Tris – 12.1gm

2.0M NaCl – 3.7gm

10mM EDTA – 116.8gm

Exercise – 2

Isolation of genomic DNA from prokaryotes

Principle The genetic material (genome) in bacteria is not very well organized as compared to eukaryotic

genomes, which is highly condensed and are present as nucleosomes .So extraction of bacterial genomic

DNA is fairly simple and do not involve elaborate protocols. Three major types of techniques or

combination of them are employed in isolation of nucleic acids: differential solubility, adsorption

methods or density gradient centrifugation. Choice of method depends on source of DNA being isolated

and its application. Major goal of nucleic acid isolation is removal of proteins, which is accomplished due

to their different chemical properties. Most nucleic acid isolation protocols involve:

a. Cell lysis step

b. Enzymatic treatments

c. Differential solubility (phenol extraction or adsorption to solid support)

d. Precipitation

Cell lysis:

Nucleic acids (DNA/RNA) must be solubilized from cells or other biological material. This solubilization usually carried out under denaturing conditions such as SDS, alkali, boiling or chaotropic agent. These denaturing conditions efficiently solubilize the nucleic acids and generally do not adversely affect them. In addition, the denaturing conditions promote the removal of proteins during the subsequent steps and inhibit the activity of nucleases that degrade the nucleic acids.

Enzymatic treatment:

Unwanted components (like proteins and RNA) are degraded by enzymes

. Proteases (Proteinase K): To remove proteins

. RNases (RNase A): To get rid of RNA

Differential solubility:

• Phenol Extraction:

Phenol is an organic solvent used to separate proteins from DNA. It is mixed in equal volumes with the DNA. The two phases are then separated by centrifugation and the upper aqueous phase that contains the nucleic acid is retained. Proteins are seen as flocculent material at the interface.

• Adsorption to solid support:

DNA is known to adsorb to silica gel surfaces in presence of high concentration of chaotropic salts that remove water from hydrated molecules in solution. Polysaccharides and proteins do not

adsorb to silica gel and are thus removed. DNA is then eluted out under low-salt conditions, free of RNA.

Precipitation:

DNA is precipitated from dilute solutions with ethanol or isopropanol, in presence of sodium or potassium acetate, pH 5.0-5.5, added to a final concentration of 0.3 M. Sodium and acidic pH will neutralize the highly charged phosphate backbone and promote hydrophobic interactions. The precipitated DNA is collected by centrifugation or is spooled out with a Pasteur pipette. The pellet is rinsed with 70% ethanol to remove any excess salt, dried and dissolved in an appropriate buffer.

Procedure:

. Take 1.5 ml of the overnight culture of bacteria and centrifuge at 6000rpm for 10 min.

. Resuspend the cells in 700ul of Cell lysis solution at RT.

. Incubate at RT for 30 minutes and spin at 10,000 rpm for 10 minutes.

. Collect 500 of the supernatant in a fresh vial, avoid taking the pellet.

. Add equal volume of phenol, chloroform, Isoamyl alcohol and mix by inverting and centrifuge at 6000 rpm for 10 min.

. Collect the upper aqueous layer in a fresh eppendorf.

. Add 1 ml of alcohol to 500 of supernatant. Mix by inverting the tube till white strands of

. DNA precipitating out is seen.

. Spin at 10,000 rpm for 10 minutes and discard the supernatant or spool precipitated DNA with the help of a tip and transfer in to a fresh vial.

. Add 0.5 ml of 70% alcohol to the DNA pellet and give a short spin at 10,000 rpm. Decant the alcohol solution taking care not to dislodge the pellet. Repeat this step once again.

. Add 100ul of T.E Buffer and incubate for 5 minutes at 50-550C to increase the solubility of Genomic DNA.

. Spin at 10,000 rpm for 10 minutes to remove insoluble material.

. Pipette out the supernatant into a fresh vial.

. Add 2.5ul of gel loading buffer to 25ul of freshly extracted DNA samples.

. Load the samples along with 10ul of control DNA on a 1% agarose gel. (Note down the order in which the samples have been loaded).

. Electrophoreses the samples at 100 volts for 1-2 hours.

. Visualize the band(s).

Observation:

Band of extracted Genomic DNA was observed and was compared with the control.

Exercise – 3

Isolation of genomic DNA from plants

Principle:

The search for a more efficient means of extracting DNA of both higher quality and yield has lead to the development of variety of protocols, however the fundamentals of DNA extraction remains the same. DNA must be purified from cellular material in a manner that prevents degradation. Because of this, even crude extraction procedures can still be adopted to prepare a sufficient amount of DNA to allow for multiple end uses.

DNA extraction from plant tissue can vary depending on the material used. Essentially any mechanical means of breaking down the cell wall and membranes to allow access to nuclear material, without its degradation is required. For this, usually an initial grinding stage with liquid nitrogen is employed to break down cell wall material and allow access to DNA while harmful cellular enzymes and chemicals remain inactivated. Once the tissue has been sufficiently ground, it can then be resuspended in a suitable buffer, such as CTAB. In order to purify DNA, insoluble particulates are removed through mixing with chloroform and centrifugation. DNA must then be precipitated from the aqueous phase and washed thoroughly to remove contaminating salts. The purified DNA is then resuspended and stored in TE buffer or sterile distilled water. This method has been shown to give intact genomic DNA from plant tissue. To check the quality of the extracted DNA, a sample is run on an agarose gel, stained with Ethidium bromide, and visualized under UV light.

Materials

• CTAB buffer

• Microfuge tubes

• Mortar and pestle

• Liquid Nitrogen

• Microfuge

• Absolute Ethanol (ice cold)

• 70% Ethanol (ice cold)

• M Ammonium Acetate

• 550C water bath

• Chloroform: Iso Amyl Alcohol (24:1)

• Water (sterile)

• Agarose

• 6X Loading Buffer

• 1X TBE solution

• Agarose gel electrophoresis system

• Ethidium Bromide solution

Preparations:

CTAB buffer 100ml

2.0 g CTAB (hexadecyl trimethyl-ammonium bromide)

10.0 ml 1M Tris pH 8.0

4.0 ml 0.5 M EDTA pH 8.0

28.0ml 5 M NaCl

40.0 ml H2O

1 g PVP 40 (polyvinyl pyrrolidine homopolymer)

Adjust all to pH 5.0 with HCl and make up to 100ml with H2O.

1M Tris pH 8.0

Dissolve 121.1g of Tris base in 800ml of H2O. Adjust pH to 8.0 by adding 42ml of concentrated HCl. Allow the solution to cool to room temperature before making the final adjustments to pH. Adjust the volume to 1L with H2O. Sterilize using an autoclave.

5x TBE buffer

54g Tris base

27.5g boric acid

20ml of 0.5M EDTA (pH 8.0)

Make up to 1L with water

To make a 0.5x working solution, do a 1:10 dilution of the concentrated stock.

1% Agarose gel

1g Agarose dissolved in 100ml TBE

Procedure

- Grind 200 mg of plant tissue to a fine paste in approximately 500 ul of CTAB buffer. - Transfer CTAB/plant extract mixture to a Microfuge tube. - Incubate the CTAB/plant extract mixture for about 15 min at 550C in a recirculating

water bath. - After incubation, spin the CTAB/plant extract mixture at 12000 g for 5 min to spin down

cell debris. Transfer the supernatant to clean Microfuge tubes. - To each tube add 250 ul of Chloroform: Iso Amyl Alcohol (24:1) and mix the solution by

inversion. After mixing, spin the tubes at 12000 rpm for 2 min. - Transfer the upper aqueous phase only (contains the DNA) to a clean Microfuge tube - To each tube add 50ul of 7.5 M Ammonium Acetate followed by 500 ul of ice cold

absolute ethanol - Invert the tubes slowly several times to precipitate the DNA. Generally the DNA can be

seen to precipitate out of solution. Alternatively the tubes can be placed for 1 hr at -200C after the addition of ethanol to precipitate the DNA.

- Following precipitation, the DNA can be pipeted off by slowly rotating/spinning a tip in the cold solution. The precipitated DNA sticks to the pipette and is visible as a clear thick precipitate. To wash the DNA, transfer the precipitate into a Microfuge tube containing 500ul of ice cold 70% ethanol and slowly invert the tube. Repeat (alternatively the precipitate can be isolated by spinning the tube at 13000 rpm for a minute to form a pellet. Remove the supernatant and wash the DNA pellet by adding two changes of ice cold 70% ethanol).

- After the wash, spin the DNA into a pellet by centrifuging at 13000 rpm 1 min. Remove all the supernatant and allow the DNA pellet to dry (approximately 15 min). Do not allow the DNA to over dry or it will be hard to re-dissolve.

- Resuspend the DNA in sterile DNase free water (approximately 50-400 ul H2O; the amount of water needed to dissolve the DNA can vary. Depending on how much is isolated). RNaseA (10ug/ml) can be added to the water prior to dissolving the DNA to remove any RNA in the preparation (10 ul RNase A in 10 ml H2O)

- After resuspension, the DNA is incubated at 650C for 20 min to destroy any DNases that may be present and store a 40C.

- Agarose gel electrophoresis of the DNA will show the integrity of the DNA, while spectrophotometry gives an indication of the concentration and cleanliness.

DNA quality confirmation - Prepare a 1% solution of agarose by melting 1g of agarose in 100 ml of 0.5x TBE buffer

in a microwave for approximately 2 min. Allow to cool for a couple of minutes then add 2.5 ul of Ethidium bromide. Stir to mix.

- Cast a gel using a supplied try and comb. Allow the gel set for a minimum of 20 min at room temperature on a flat surface.

- Load the following in to separate wells

• 10 ul. 1kb ladder

• 5 ul sample+ 5uL water+2 ul 6x loading Buffer - Run the gel for 30 min at 100 V - Expose the gel to UV light and photograph (demonstration) - Confirm DNA quality, presence of a highly resolve high molecular weight ban indicates

good quality DNA. Presence of a smeared band indicates DNA degradation.

Observation:

Band of extracted Genomic DNA was observed and was compared with the control. Also band of RNA was observed.

Exercise – 4

Qualitative and Quantitative analysis of Nucleic acid

Principle:

After isolation of DNA, quantitative and qualitative analysis of nucleic acid is necessary to ascertain the approx. quantity of DNA obtained and the suitability of DNA for further analysis. This is important for many applications including digestion of DNA by restriction enzyme or PCR amplification of target DNA.

The most common methodology for quantifying the amount of nucleic acid in preparation is spectrophotometric analysis. If sample amount is less, the former method is most suitable.

Spectrophotometric analysis:

Analysis of UV absorption by the nucleotides provides a simple and accurate estimation of concentration of nucleic acid in a sample. Purines and pyrimidines in nucleic acid show absorption maxima around 260nm (e.g. dATP: 259nm, dCTP: 272nm, dTTP: 247nm). If the DNA sample is pure without significant contamination from protein or organic solvents, the ratio of OD260/OD 280 should be determined to asses the sample purity. This method is however limited by quantity of DNA and purity of preparation. Accurate analysis of DNA preparation may be impeded by the presence of impurity on the sample or very less amount of DNA. Also presence of RNA, sheared DNA etc. could interfere with the accurate estimation of total high molecular weight genomic DNA.

• According to Beer – Lambert law absorbance is directly proportional to the concentration and is inversely proportional to the path length.

• The amount of DNA can be calculated quantitatively by use of formula Yield = OD260* 50*dilution factor/1000ug/ml

• The purity or quality of the extracted DNA can be calculated by use of formula Purity = OD260/OD 280

Protocol:

1. Take 1ml of TE buffer in a quartz cuvette and calibrate spectrophotometer. 2. Add 100ul of each sample to 900ul of TE buffer and mix well. 3. Use TE buffer as blank. 4. Note the OD260 and OD 280 values on spectrophotometer. 5. Calculate the ratio of OD260/OD 280.

Observation:

Spectrophotometric conversion of nucleic acid

• Absorbance 1 at 260nm represents 50ug/ml of ds DNA. • Absorbance 1 at 260nm represents 33ug/ml of ss oligonucleotides.

• Absorbance 1 at 260nm represents 40ug/ml of ss RNA.

• An A260/A280 ratio between 1.7 and 2.0 generally represents a high quality DNA sample.

Observation table:

sample OD260 OD280 Sample 1

0.086

0.093

Sample 2

0.058

0.056

Sample 3

0.045

0.048

Sample 4

0.073

0.084

Sample Purity of DNA (A260/A280)

Conclusion

Sample 1

0.086/0.093 = 0.92

contaminate

Sample 2

0.058/0.056 = 1.035

contaminate

Sample 3

0.045/0.048 = 0.9375

contaminate

Sample 4

0.073/0.084 = 0.8690

contaminate

Calculation of amount of DNA at A260

Sample 1

= OD260* 50*dilution factor/1000ug/ml

= 0.086* 50*10/1000ug/ml

= 0.043ug/ml

Sample 2

= OD260* 50*dilution factor/1000ug/ml

= 0.058* 50*10/1000ug/ml

= 0.029ug/ml

Sample 3

= OD260* 50*dilution factor/1000ug/ml

= 0.045* 50*10/1000ug/ml

= 0.0225ug/ml

Sample 4

= OD260* 50*dilution factor/1000ug/ml

= 0.073* 50*10/1000ug/ml

= 0.0365ug/ml

Exercise – 5

Determination of melting temperature of DNA

Principle:

The melting temperature, Tm, of an oligonucleotide is its most critically important value. The most reliable and accurate determination melting temperature is determined empirically. The main factors affecting Tm are salt concentration, strand concentration, and the presence of denaturants (such as formamide or DMSO). Other effects such as sequence, length, and hybridization conditions can be important as well. Tm – The temperature at which 50% of the oligonucleotide and its perfect complement are in duplex. Single strand DNA absorbs more UV light than double strands. Moreover, ,double strands can be separated by heat (melted) and the temperature at which the strands separate (T m) is related to the number of guanine-cytosine residues( each having three hydrogen bonds as opposed to the two in adenine- thymine). This has led to the development of rapid test for an approximation of the GC/AT ratio using melting points and the change in UV absorbance (known as “hyperchromicity” or “hyperchromatic shift”). Of course, the separation is also dependent upon environmental influences, particularly the salt concentration of the DNA solution. To standardize this, all TM measurements are made in TE buffer. DNA melts between 850 and 1000C in this buffer.

Materials

DNA

T E Buffer

UV spectrophotometer (preferably with temperature control)

Procedure

. Place the cuvettes into a waterbath at 250C and allow to temperature equilibrate.

. Remove the blank, wipe the outside dry and rapidly blank the instrument at 260 nm. Transfer the sample to the spectrophotometer (be sure to dry and work rapidly) and read the absorbance.

. Raise the temperature of the bath to 500C and repeat step a.

. Raise the temperature sequentially to 60oC, 650C, 700C, 750C and 800C and repeat the absorbance measurements.

. Slowly raise the temperature above 800 and make absorbance measurements every 20 until the absorbance begins to increase. At that point, increase the temperature, but continue to take readings at 10C intervals.

. Plot the value of at vs. temperature and calculate the midpoint of any increased absorbance. This midpoint is the melting point (Tm) for your DNA sample.

Temperature A260

400C 0.392

450C 0.394

500C 0.412

550C 0.434

600C 0.457

650C 0.476

700C 0.496

750C 0.523

800C 0.548

850C 0.618

900C 0.662

950C 0.688

1000C 0.735

1050C 0.800

1100C 0.482

Observation: The melting temperature of DNA was found to be 450C.

Exercise – 6

Sodium Dodecyl Sulphate PolyAacryalmide Gel Electrophoresis

Principle:

SDS-PAGE is the most widely used method for qualitatively analyzing any protein mixture, monitoring protein purity and to determine their molecular weights. It is based on the separation of proteins according to their size and then locating them by binding to a dye. SDS or sodium Dodecyl sulphate is an anionic detergent that binds strongly to proteins, causing their denaturation. In the presence of excess SDS, about 1.4 g of the detergent binds to each gram of protein, giving the [protein a constant negative charge per unit mass. As a result, protein-SD complex move towards the anode during electrophoresis and owing to molecular sieving properties of the polyacrylamide gel, protein get separated based on their molecular weights. Since, the principle of this technique is separation of proteins based on size differences, by running standard proteins of known molecular weights on the same gel as unknown protein, molecular weight of the unknown protein can be determined. Mobility of protein in SDS gel electrophoresis is expressed as relative mobility (RF) with respect to the tracking dye, bromo phenol blue.

RF = distance migrated by protein/distance migrated by tracking dye

RF value of protein marker of known size is used to generate a standard curve by plotting the molecular weights against the RF value on semi-log graph. The molecular weight of the unknown protein can then be extrapolated from its RF value.

The technique consists of two basic steps:

Step 1: preparation of polyacrylamide Gel

Cross linked polyacrylamide gels are formed by co-polymerization of acrylamide monomer and a cross-linking agent N, N’ – Methylene bisacrylamide. This reaction is catalyze by N, N, N’, N’- Tetramethyenediamine (TEMED) and initiated by ammonium persulphate (APS) Porosity of the gel is determined by the amount of acrylamide and bis-acrylamide mix used. Lower percentage gels have larger pore sizes, thereby offering less resistance to passage of larger molecules, while higher percentage gels favour separation smaller molecules. Most protein separations are carried out using gels ranging from 5-15% acrylamide.

Step 2: Electrophoresis

The polyacrylamide gel slab is prepared and fixed to a vertical electrophoresis apparatus. Protein samples are usually denatured by boiling in sample loading buffer containing Tris buffer of pH 6.8, SDS,2 Beta Mercaptoethanol (reduces disulphide bonds), sucrose glycerol (to increase the density), and bromo phenol blue ( As a tracking dye) and loaded onto wells in the gel.

Procedure:

1. Assemble the plates for casting gel. 2. Clamp the assembly of plates. Ensure the assembly is leak proof by filling water between the plates.

Silicon grease can be applied to spacers or 1% agarose can be used for sealing to make it leak proof. 3. Prepare the Resolving gel by mixing the individual component in the fixed required volume.

Reagent 12.5% to make 30ml Distilled water 9.6ml (30%/0.8%) Acrylamide/Bisacrylamide 12.5ml Tris HCl buffer(1.5M pH 8.8) 7.5ml 10% SDS 300ul Ammonium per sulphate 150ul TEMED 20ul

4. Pour the gel solution between the plates till the level is about 2 cm below the top edge of notched plate.

5. Add 200 to 250ul of water to make the surface even. 6. After the gel are set (approximately 20-30 minutes), wash the top of the separating gel with

distilled water and drain off the water completely. 7. Prepare the Stacking gel by mixing the individual component in the fixed required volume.

Reagent 5% to make 15ml Distilled water 10.3ml (30%/0.8%) Acrylamide/Bisacrylamide 2.49ml Tris HCl buffer(1.5M pH 8.8) 1.89ml 10% SDS 150ul Ammonium per sulphate 150ul TEMED 15ul

8. Insert the comb into the gel solution carefully without trapping any air bubbles, approximately 1 cm above the separating gel. The stacking gel will set in about 10 minutes.

9. Pipette out 25ul of protein samples and 10 of protein marker into individual vials. Label them appropriately. To each of these vials add 15 of sample loading buffer.

Reagent 5% to make 15ml 10% SDS 4ml 10mM beta mercapto ethanol 0.8ml 20% glycerol 2ml 0.2M Tris HCl buffer(pH 6.8) 2.5ml 0.05% bromophenol blue 300ul

10. Place the vials in a boiling water bath for 5 minutes. 11. After stacking gel has set, carefully remove the comb and the bottom spacer. Wash the wells

immediately with distilled water to remove non-polymerized acrylamide. 12. Fill the bottom reservoir with 1X reservoir buffer.

13. Carefully fix the plate to PAGE apparatus without trapping any air bubbles between the buffer and the bottom of the gel, with the noticed plate facing the top reservoir.

14. Fill the top reservoir with the 1X reservoir buffer. 15. Load samples in to the wells; rinse the micropipette tip in the bottom reservoir buffer between

each load. Note down the order in which the samples have been loaded. 16. Connect the cords to the power supply, according to the convention.

Red: anode, black: cathode

17. Set voltage at 100 V and switch on the power supply. 18. When the dye front reaches 0.5 cm above the bottom of the gel, turn off the power. 19. Remove the gel plates and gently pry the plates apart. Use a spatula or similar tool to separate the

plates (not at the notch). 20. Transfer the gel to a tray containing water; wash the gel for 5 minutes. Discard the water. 21. Add 20 ml of Staining Solution and stain the gel for 30-60 minutes. 22. Destain the gel with distaining solution. 23. Destain the gel with water, if the background is not clear. 24. Use a ruler to measure the distance migrated by each band of the protein marker from the start of

separating gel. 25. Similarly, measure the distance migration distance of protein sample C and the tracking dye from

the start of separating gel.

Observation:

Distinct band of protein was obtained as the protein taken as sample was BSA fraction. Multiple bands of protein were observed as compared to control.

Dark blue bands were obtained on light blue background of gel.

Exercise – 7

Polymerase Chain Reaction technique

Principle Developed in 1983 by Kary Mullis, PCR is now a common and often indispensable technique.

The polymerase chain reaction (PCR) is a technique in molecular biology to amplify a single or few copies

of a piece of DNA across several orders of magnitude, generating thousands to millions of copies of a

particular DNA sequence. The method relies on thermal cycling, consisting of cycles of repeated heating

and cooling of the reaction for DNA melting and enzymatic replication of the DNA. Primers (short DNA

fragments) containing sequences complementary to the target region along with a DNA

polymerase (after which the method is named) are key components to enable selective and repeated

amplification. As PCR progresses, the DNA generated is itself used as a template for replication, setting

in motion a chain reaction in which the DNA template is exponentially amplified. PCR can be extensively

modified to perform a wide array of genetic manipulations.

Almost all PCR applications employ a heat-stable DNA polymerase, such as Taq polymerase, an enzyme

originally isolated from the bacterium Thermus aquaticus. This DNA

polymerase enzymatically assembles a new DNA strand from DNA building blocks, the nucleotides, by

using single-stranded DNA as a template and DNA oligonucleotides (also called DNA primers), which are

required for initiation of DNA synthesis. The vast majority of PCR methods use thermal cycling, i.e.,

alternately heating and cooling the PCR sample to a defined series of temperature steps. These thermal

cycling steps are necessary first to physically separate the two strands in a DNA double helix at a high

temperature in a process called DNA melting. At a lower temperature, each strand is then used as

the template in DNA synthesis by the DNA polymerase to selectively amplify the target DNA. The

selectivity of PCR results from the use of primers that are complementary to the DNA region targeted

for amplification under specific thermal cycling conditions.

Extraction of DNA from the provided Sample

Requirements:

Chemicals and Reagents:

1. Sodium hydroxide 2. Sodium citrate 3. Proteinase K 4. Phosphate buffer

Instruments:

1. Laminar Air flow 2. Dry Bath 3. Centrifuge

Protocol:

1. Take Equal volume of 0.1 M Sodium citrate and 1 M NaOH. 2. Mix Equal Volume of sample with it. 3. Incubate the Mix for 20 min. 4. Centrifuge the Mix at 6000 rpm for 20 min. 5. Decant supernatant and add 1 ml of phosphate buffer. 6. Centrifuge at 6000 rpm for 5 min. 7. Decent supernatant and add 1 ml of phosphate buffer. 8. Centrifuge at 2500 rpm for 2-3 min. 9. Take 1 ml of supernatant in an eppendorf tube & centrifuge at 10000 rpm for 5 min. 10. To the pellet add 90 ul of TE buffer & 10 ul of Proteinase K. 11. Vortex to Mix and Incubate at 600C for 3 hrs after that at 1000C for 10 min. 12. Keep at 40 for 5 min and centrifuge at 10000 rpm for 2 min. 13. Supernatant hence obtained contains extracted DNA, which is then amplified using

thermal cycler.

Amplification of the extracted DNA

Requirements:

Chemicals and Reagents

1. PCR Buffer 2. Magnesium chloride solution 3. dNTP’s solution 4. Primer 5. Nuclease free water 6. Taq Polymerase

7. Template DNA

Instrument:

1. Thermal cycler 2. Laminar Air Flow

Protocol:

1. All the reagents are taken out from -200C and thawed. 2. The Pre PCR mix is prepared from the solutions by adding the individual component in

required amount so that the desired concentration is attained. 3. Template DNA is added in required amount.

Composition of PCR Mix:

o PCR buffer 2.5 ul o MgCl2 2.5 ul o d NTP’s 2ul o Primer 1 1ul o Primer 2 1ul o Taq polymerase 0.1ul o Distilled Water 10.9ul o Template DNA 5ul o The pre PCR mix hence prepared is placed in thermal cycler and Allowed to

amplify by carrying out 30-35 cycles. o After completion of the amplification cycle the amplified product is taken out

from the Thermal cycler and run on an Agarose gel to visualize the results.

Electrophoresis of the Amplified DNA Product

Requirements:

Chemicals and Reagents

1. Agarose 2. 50X TE Buffer

3. Distilled water 4. Ethidium bromide

Instruments:

1. Electrophoresis unit 2. Power pack 3. UV Transilluminator

Protocol:

1. Prepare Ix TAE by diluting appropriate amount of 50X TAE buffer. 2. Weight 0.5 g of agarose and add to 50 ml of ix TAE. This gives 1% agarose gel. 3. Boil till agarose dissolves completely and a clear solution results. 4. Meanwhile place the combs of electrophoresis set such that it is approximately 2 cm

away from the cathode. 5. Pour the agarose solution in the central part of tank when the temperature reaches

approximately 600C do not generate air bubbles. The thickness of the gel should be around 0.5 to 0.9 cm. Keep the gel undisturbed at room temperature for the agarose to solidify.

6. Pour IX TAE buffer in to the gel tank till the buffer level stands at 0.5 to 0.8 cm above the gel surface.

7. Gently lift the combs, ensuring that wells remain intact. 8. Connect the power cord to the electrophoresis power supply according to the

convention Red: anode, black: cathode.

9. Load the samples in the wells in the desired order. 10. Set the voltage to 50 V and switch on the power supply. 11. Allow the gel to run for more then half the length of gel. 12. Switch off the power supply and visualized Gel under UV- Transilluminator for

interpretation of the result.

Observation: Positive result was interpreted as a distinct band of DNA was observed as compared to the control of

123bp sequence IS6110 specific to Mycobacterium tuberculosis.