NEHRU ARTS AND SCIENCE COLLEGE

DEPARTMENT OF MICROBIOLOGY

E-LEARNING

CLASS : II B.Sc. Microbiology

SUBJECT : BIOINSTRUMENTATION-PRINCIPLES AND APPLICATIONS

SEMESTER –III

UNIT – I

Microscopy– Principles and application – Bright field, Darkfield, Phase contrast, Fluorescence,

SEM & TEMS- Specimen preparation of Electron microscopy .

UNIT – II

Principles and Applications of Autoclave , Hot air oven , Incubator , Laminar air flow, BOD

incubator, Metabolic shaker , Incinerator.

UNIT -III

Centrifuges –Low speed, High speed , Ultra centrifuge. pH meter , Lyophilizer.

UNIT –IV

Colorimetry, Turbidometry, Spectrometry – UV & Visible Spectrophotometer . Flame

Photometry.

UNIT-V

Chromatography – Paper , Thinlayer, Column, Ion-exchange, Gas and HPLC .

Electrophoresis – SDS – PAGE and Agarose gel electrophoresis.

PART-A

1.What magnification is used if you observe a microorganism with a microscope whose object is 100× and whose ocular lens is 10×?(a) 1000× magnification(b) 100× magnification(c) 10× magnification(d) 10,000× magnification2.What is the function of an illuminator?(a) To control the temperature of the specimen(b) To keep the specimen moist(c) An illuminator is the light source used to observe a specimen under a microscope(d) To keep the specimen dry3.What is the area seen through the ocular eyepiece called?(a) The stage(b) The objective(c) The display(d) The field of view4.How do you maintain good resolution of a specimen at magnifications greater than 100?(a) Display the specimen on a television monitor.(b) Use a single ocular eyepiece.(c) Immerse the specimen in oil.(d) Avoid moving the specimen.5.What is a micrograph?(a) A microscopic photograph taken by an electron microscope(b) A microscopic diagram of a specimen(c) A microscopic photograph taken by a light microscope(d) A growth diagram of a specimen6.What is refractive index

Light waves that are reflected by the specimen are measured by the refractive index. The refractive index specifies the amount of light waves that is reflected by an object. There is a low contrast between a specimen and the field of view if they have nearly the same refractive index. The further these refractive indexes are from each other, the greater the contrast between the specimen and the field of view.7. Sterilization

Sterilization is the destruction of all microorganisms and viruses, as well as endospores. Sterilization is used in preparing cultured media and canned foods. It is usually performed by steam under pressure, incineration, or a sterilizing gas such as ethylene oxide.8.Antisepsis

Antisepsis is the reduction of pathogenic microorganisms and viruses on living tissue. Treatment is by chemical antimicrobials, like iodine and alcohol. Antisepsis is used to disinfect living tissues without harming them.9.Commercial sterilization

Commercial sterilization is the treatment to kill endospores in commercially canned products. An example is the bacteria Clostridium botulinum, which causes botulism.

10.AsepticAseptic means to be free of pathogenic contaminants. Examples include proper hand

washing, flame sterilization of equipment, and preparing surgical environments and instruments.11.Disinfection

Disinfection is the destruction or killing of microorganisms and viruses on nonliving tissue by the use of chemical or physical agents. Examples of these chemical agents are phenols, alcohols, aldehydes, and surfactants.12.Degerming

Degerming is the removal of microorganisms by mechanical means, such as cleaning the site of an injection. This area of the skin is degermed by using an alcohol wipe or a piece of cotton swab soaked with alcohol. Hand washing also removes microorganisms by chemical means.13.Pasteurization

Pasteurizationuses heat to kill pathogens and reducethe number of food spoilage microorganisms in foods and beverages. Examples are pasteurized milk and juice.14.Sanitation

Sanitation is the treatment to remove or lower microbial counts on objects such as eating and drinking utensils to meet public health standards. This is usually accomplished by washing the utensils in high temperatures or scalding water and disinfectant baths. acterostatic, fungistatic, and virustastic agents— or any word with the suffix -static or -stasis—indicate the inhibition of a particular type of microorganism. These are unlike bactericides or fungicides that kill or destroy the organism. Germistatic agents include refrigeration, freezing, and some chemicals.15.What is meant by buffer solution?

Buffers are solutions, which could tolerate small changes in pH during reactions.

The buffer is usually composed of a weak acid and its salts. It is indispensable for performing

biological reactions at their optimal rate in vitro. For example, a solution of acetic acid and its

salt, sodium acetate, form an effective buffer system.

Buffers play an important role in vivo also, because most biological reactions require

optimal pH. Various substances like phosphates, carbonates, amino acids and proteins in cells

maintains a constant pH. Therefore in vitro studies of biological reaction require buffers.

PART B1. Describe the prefixes for the metric system and their equivalent in meters.

Prefix Value in MetersKilo (km) (kilo = 1,000) 1,000 mDeci (dm) (deci = 1/10) 0.10 mCenti (cm) (centi = 1/100) 0.01 mMilli (mm) (milli = 1/1000) 0.001 mNano (nm) (nano = 1/1,000,000,000) 0.000000001 mPico (pm) (pico = 1/1,000,000,000,000) 0.000000000001 mKilo (kg) 1,000 gHecto (hg) 100 gDeka (dag) 10 gGram (g) 1 gDeci (dg) 0.1 gCenti (cg) 0.01 gMilli (mg) 0.001 gMicro (μg) 0.000001 gNano (ng) 0.000000001 gPico (pg) 0.000000000001 g

Ameter is the standard for length in the metric system. Akilogram is the standard for mass in the metric system. A gram uses the same prefixes as a meter to specify the number of grams that are represented by a value. For example, a kilometer is 1,000 meters and a kilogram is 1,000 grams. This makes it a lot easier to learn the metric system since the number of grams and meters are indicated by the same set of prefixes. 2.Write about various parts of compound microscope

A microscope is a complex magnifying glass. In the 1600s, during the time of Antoni van Leeuwenhoek, microscopes consisted of one lens that was shaped so that the refracted light magnified a specimen 100 times its natural size. Other lenses were shaped to increase the magnification to 300 times. However, van Leeuwenhoek realized that a single-lens microscope is difficult to focus. Once Van Leeuwenhoek brought the specimen into focus, he kept his hands behind his back to avoid touching the microscope for fear they would bring the microscope out of focus. It was common in the 1600s for scientists to make a new microscope for each specimen that wanted to study rather than try to focus the microscope. The single-lens magnifying lens or glass is a thing of the past. Scientists today use a microscope that has two sets of lenses (objective and ocular), which is called a compound light microscope. Fig. shows parts of a compound light microscope. A compound light microscope consists of:• Illuminator. This is the light source located below the specimen.• Condenser. Focuses the light through the specimen.• Stage. The platform that holds the specimen.• Objective. The lens that is directly above the stage.• Nosepiece. The portion of the body that holds the objectives over the stage.• Field diaphragm. Controls the amount of light into the condenser.• Base. Bottom of the microscope.

•Coarse focusing knob. Used to make relatively wide focusing adjustments to the microscope.• Fine focusing knob. Used to make relatively small adjustments to the microscope.• Body. The microscope body.• Ocular eyepiece. Lens on the top of the body tube. It has a magnification of 10× normal vision.

Parts of a compound light microscope.

3.How to measure magnification in the microscope? Explain.A compound microscope has two sets of lenses and uses light as the source of

illumination. The light source is called an illuminator and passes light through a condenser and through the specimen. Reflected light from the specimen is detected by the objective. The objective is designed to redirect the light waves, resulting in the magnification of the specimen.

There are typically four objectives, each having a different magnification. These are 4×, 10×, 40×, and 100×. The number indicates by how many times the original size of a specimen is magnified, so the 4× objective magnifies the specimen four times the specimen size. The eyepiece of the microscope is called the ocular eyepiece and it, too, has a lens—called an ocular lens—that has a magnification of 10×.

You determine the magnification used to observe a specimen under a microscope by multiplying the magnification of the objective by the magnification of the ocular lens. Suppose you use the 4× objective to view a specimen. The image you see through the ocular is 40× because the magnification of the object is multiplied by the magnification of the ocular lens, which is 10×.

4.What is resolution? Explain about it.The area that you see through the ocular eyepiece is called the field of view. Depending

on the total magnification and the size of the specimen, sometimes the entire field of view is filled with the image of the specimen. Other times, only a portion of the field of view contains the image of the specimen.

You probably noticed that the specimen becomes blurry as you increase magnification. Here’s what happens. The size of the field of view decreases as magnification increases, resulting in your seeing a smaller area of the specimen. However, the resolution of the image remains unchanged, therefore you must adjust the fine focus knob to bring the image into focus again.

Resolution is the ability of the lens to distinguish fine detail of the specimen and is determined by the wavelength of light from the illuminator. At the beginning of this chapter you learned about the wave cycle, which is the process of the wave going up and then falling down time and again. A wavelength is the distance between the peaks of two waves. As a general rule, shorter wavelengths produce higher resolutions of the image seen through the microscope.5.How to maintain good resolution at magnifications of 100× and greater? Explain

In order to maintain good resolution, the lens must be small and sufficient light must be reflected from both the specimen and the stain used on the specimen. The problem is that too much light is lost; air between the slide and the objective prevents some light waves from passing to the objective, causing the fuzzy appearance of the specimen in the ocular eyepiece.

The solution is to immerse the specimen in oil. The oil takes the place of air and, since oil has the same refractive index as glass, the oil becomes part of the optics of the microscope. Light that is usually lost because of the air is no longer lost. The result is good resolution under high magnification.6.Write about different types of specimen preparation

There are two ways to prepare a specimen to be observed under a light compoundmicroscope. These are a smear and a wet mount.SmearA smear is a preparation process where a specimen that is spread on a slide. Smear can prepare using the heat fixation process:

1. Use a clean glass slide.2. Take a loop of the culture.3. Place the live microorganism on the glass slide.4. The slice is air dried then passed over a Bunsen burner about three times.5. The heat causes the microorganism to adhere to the glass slide. This is known as fixing the microorganism to the glass slide.6. Stain the microorganism with an appropriate stain.

Wet MountA wet mount is a preparation process where a live specimen in culture fluid is placed on a

concave glass side or a plain glass slide. The concave portion of the glass slide forms a cup-like shape that is filled with a thick, syrupy substance, such as carboxymethyl cellulose. The microorganism is free to move about within the fluid, although the viscosity of the substance slows its movement. This makes it easier for you to observe the microorganism. The specimen and the substance are protected from spillage and outside contaminates by a glass cover that is placed over the concave portion of the slide.

7.What is stain? Explain about itA stain is a chemical that adheres to structures of the microorganism and in effect dyes

the microorganism so the microorganism can be easily seen under a microscope. Stains used in microbiology are either basic or acidic. Basic stains are cationic and have positive charge.

Common basic stains are methylene blue, crystal violet, safranin, and malachite green. These are ideal for staining chromosomes and the cell membranes of many bacteria.

Acid stains are anionic and have a negative charge. Common acidic stains are eosin and picric acid. Acidic stains are used to stain cytoplasmic material and organelles or inclusions.

8.Describe how an autoclave works. What conditions are required for sterilization by moist heat, and what three things must one do when operating an autoclave to help ensure success?

Moist heat sterilization must be carried out at temperatures above 100°C in order to destroy bacterial endospores, and this requires the use of saturated steam under pressure. Steam sterilization is carried out with an autoclave (figure), a device somewhat like a fancy pressure cooker. The development of the autoclave by Chamberland in 1884 tremendously stimulated the growth of microbiology. Water is boiled to produce steam, which is released through the jacket and into the autoclave’s chamber. The air initially present in the chamber is forced out until the chamber is filled with saturated steam and the outlets are closed. Hot, saturated steam continues to enter until the chamber reaches the desired temperature and pressure, usually 121°C and 15

pounds of pressure. At this temperature saturated steam destroys all vegetative cells and endospores in a small volume of liquid within 10 to 12 minutes. Treatment is continued for about 15 minutes to provide a margin of safety. Of course, larger containers of liquid such as flasks and carboys will require much longer treatment times.

Moist heat is thought to kill so effectively by degrading nucleic acids and by denaturing enzymes and other essential proteins. It also may disrupt cell membranes.

Autoclaving must be carried out properly or the processed materials will not be sterile. If all air has not been flushed out of the chamber, it will not reach 121°C even though it may reach a pressure of 15 pounds. The chamber should not be packed too tightly because the steam needs to circulate freely and contact everything in the autoclave. Bacterial endospores will be killed only if they are kept at 121°C for 10 to 12 minutes. When a large volume of liquid must be sterilized, an extended sterilization time will be needed because it will take longer for the center of the liquid to reach 121°C; 5 liters of liquid may require about 70 minutes. In view of these potential difficulties, a biological indicator is often autoclaved along with other material. This indicator commonly consists of a culture tube containing a sterile ampule of medium and a paper strip covered with spores of Bacillus stearothermophilus or Clostridium PA3679. After autoclaving, the ampule is aseptically broken and the culture incubated for several days. If the test bacterium does not grow in the medium, the sterilization run has been successful. Sometimes either special tape that spells out the word sterile or a paper indicator strip that changes color upon sufficient heating is autoclaved with a load of material. If the word appears on the tape or if the color changes after autoclaving, the material is supposed to be sterile. These approaches are convenient and save time but are not as reliable as the use of bacterial endospores.

9.Explain the Law of mass action and dissociation constant of weak acid

To understand the Henderson and Hasselbalch equation, it is important to know about

dissociation constant of weak acid. Generally weak acids dissociate partially and the

concentration of H+ ions in a weak acid depends mainly on dissociation constant.

According to Law of Mass Action, rate of any reaction is directly proportional to the

concentration of the reactants. For example, the dissociation of a weak acid such as acetic acid

can be written as,

v1

CH3COOH CH3COO- + H+

v2

In such reversible reactions, the velocity of forward reaction v1 is directly proportional to

the concentration of the reactants viz, CH3COOH.

In other words, v1 α [CH3COOH] or v1 = k1 [CH3COOH]

In the same way, the velocity of the backward reaction v2, is directly proportional to the

concentration of the reactants viz, CH3COO- and H+.

v2 α [CH3COO-] [H+] or v2 = k2 [CH3COO-] [H+]

1

The k1 and k2 are defined as rate constants or proportionally constant.

The velocity of forward reaction (v1) = velocity of the backward reaction (v2). That is, at

this stage there will be no net change in the concentration of the individual species. In other

words,

k1 [CH3COOH] = k2 [CH3COO-] [H+]

On rearranging the formula , k1 [CH3COO-] [H+]

=

k2 [CH3COOH]

Since both k1 and k2 are constants, the ratio of the two will gain be a constant which is called

equilibrium constant (keq). Hence the above equation can be written as

[CH3COO-] [H+]

keq =

[CH3COOH]

Under a set of physical condition, keq remains constant and it has a great significance,

because if there is any alteration in the concentration of the any one of the components, the other

components will readjust their concentration automatically to maintain the value of keq as a

constant. The equilibrium constant also known as dissociation constant, ka.

10. Derive Henderson and Hasselbalch equation.

Henderson and Hasselbalch have extended the above equation into a useful expression

known as the Henderson – Hasselbalch equation.

In the above discussion,

1

[CH3COO-] [H+]

keq =

[CH3COOH]

In general, the above equation can be written as

[A-] [H+]

ka = or ka [AH] = [A-] [H+]

[AH]

Rearranging the terms,

Ka [AH]

[H+] =

[A-]

Taking logarithms of both sides

[AH]

log[H+] = log Ka + log

[A-]

Multiply by -1

[AH]

-log[H+] = -log Ka - log

[A-]

Since –log [H+] = pH and –log ka = pka, the above equation can be written as,

[AH] [A-]

pH = pka - log or pH = pka + log or

[A-] [AH]

[Salt]

pH = pka + log

[Acid]

This is known as Henderson – Hasselbalch equation. Note that the dissociation of weak acid

increases as the solution is diluted.

Therefore, the term [A-] increases on dilution, resulting in increase in pH.

[Ah]

11.What is Paper chromatography?

Paper chromatography is an analytical technique for separating and identifying mixtures that are or can be colored, especially pigments. This can also be used in secondary or primary colors in ink experiments. This method has been largely replaced by thin layer chromatography, however it is still a powerful teaching tool. Two-way paper chromatography, also called two-dimensional chromatography, involves using two solvents and rotating the paper 90° in between. This is useful for separating complex mixtures of similar compounds, for example, amino acids

Ascending Chromatography

In this method, the solvent is in pool at the bottom of the vessel in which the paper is supported.It rises up the paper by capillary action against the force of gravity.

Descending Chromatography

In this method, the solvent is kept in a trough at the top of the chamber and is allowed to flow down the paper. The liquid moves down by capillary action as well as by the gravitational force. In this case, the flow is more rapid as compared to the ascending method. Because of this rapid speed, the chromatography is completed in a comparatively shorter time. The apparatus needed for this case is more sophisticated. The developing solvent is placed in a trough at the top which is usually made up of an inert material. The paper is then suspended in the solvent. Substances that cannot be separated by ascending method, can be separated by the above descending method.

Rƒ value

Rƒ value may be defined as the ratio of the distance travelled by the substance to the distance travelled by the solvent. Rƒ values are usually expressed as a fraction of two decimal places but it was suggested by Smith that a percentage figure should be used instead. If Rƒ value of a solution is zero, the solute remains in the stationary phase and thus it is immobile. If Rƒ value = 1 then the solute has no affinity for the stationary phase and travels with the solvent front.

PART C1. Explain in detailed about the features of various types of microscopes

Bright-Field MicroscopeThe bright-field microscope is the most commonly used microscope and consists of two

lenses. These are the ocular eyepiece and the objective. Light coming from the illuminator passes through the specimen. The specimen absorbs some light waves and passes along other light waves into the lens of the microscope, causing a contrast between the specimen and other objects in the field of view. Specimens that have pigments contrast with objects in the field of view and can be seen by using the bright-field microscope. Specimens with few or no pigments have a low contrast and cannot be seen with the bright-field microscope. Some bacteria have low contrast.Dark Field Microscope

The dark-field microscope focuses the light from the illuminator onto the top of the specimen rather than from behind the specimen. The specimen absorbs some light waves and reflects other light waves into the lens of the microscope. The field of view remains dark while the specimen is illuminated, providing a stark contrast between the field of view and the specimen.Phase-Contrast Microscope

The phase-contrast microscope bends light that passes through the specimen so that it contrasts with the surrounding medium. Bending the light is called moving the light out of phase. Since the phase-contrast microscope compensates for the refractive properties of the specimen, you don’t need to stain the specimen to enhance the contrast of the specimen with the field of view. This microscope is ideal for observing living microorganisms that are prepared in wet mounted slides so you can study a living microorganism.Fluorescent Microscope

Fluorescent microscopy uses ultraviolet light to illuminate specimens. Some organism fluoresce naturally, that is, give off light of a certain color when exposed to the light of different color. Organisms that don’t fluoresce naturally can be stained with fluorochrome dyes. When these organisms are placed under a fluorescent microscope with an ultraviolet light, they appear very bright in front of a dark background.

Differential Interface Contrast Microscope (Nomanski)The differential interface contrast microscope, commonly known as Nomanski, works in

a similar way to the phase-contrast microscope. However, unlike the phase-contrast microscope (which produces a two-dimensional image of the specimen), the differential interface contrast microscope shows the specimen in three dimensions.

THE ELECTRON MICROSCOPEA light compound microscope is a good tool for observing many kinds of

microorganisms. However, it isn’t capable of seeing the internal structure of a microorganism nor can it be used to observe a virus. These are too small to effectively reflect visible light sufficient to be seen under a light compound microscope. In order to view internal structures of viruses and internal structures of microorganisms, microbiologists use an electron microscope where specimens are viewed in a vacuum. Developed in the 1930s, the electron microscope uses beams of electrons and magnetic lenses rather than light waves and optical lenses to view a

specimen. Very thin slices of the specimen are cut so that the internal structures can be viewed. Microscopic photographs called micrographs are taken of the specimen and viewed on a video screen. Specimens can be viewed up to 200,000 times normal vision. However, living specimens cannot be viewed because the specimen must be sliced.Transmission Electron Microscope

The transmission electron microscope (TEM) has a total magnification of up to 200,000× and a resolution as fine as seven nanometers. A nanometer is 1/1,000,000,000 of a meter. The transmission electron microscope generates an image of the specimen two ways. First, the image is displayed on a screen similar to that of a computer monitor. The image can also be displayed in the form of an electron micrograph, which is similar to a photograph. Specimens viewed by the transmission electron microscope must be cut into very thin slices, otherwise the microscope does not adequately depict the image.Scanning Electron Microscope

The scanning electron microscope (SEM) is less refined than the transmission electron microscope. It can provide total magnification up to 10,000× and a resolution as close as 20 nanometers. However, a scanning electron microscope produces three-dimensional images of specimen. The specimen must be freeze dried and coated with a thin layer of gold, palladium, or other heavy metal.

2.What are depth filters and membrane filters, and how are they used to sterilize liquids? Describe the operation of a biological safety cabinet.

Filtration is an excellent way to reduce the microbial population in solutions of heat-sensitive material, and sometimes it can be used to sterilize solutions. Rather than directly destroying contaminating microorganisms, the filter simply removes them. There are two types of filters. Depth filters consist of fibrous or granular materials that have been bonded into a thick layer filled with twisting channels of small diameter. The solution containing microorganisms is sucked through this layer under vacuum, and microbial cells are removed by physical screening or entrapment and also by adsorption to the surface of the filter material.

Depth filters are made of diatomaceous earth (Berkefield filters), unglazed porcelain (Chamberlain filters), asbestos, or other similar materials.

Membrane filters have replaced depth filters for many purposes. These circular filters are porous membranes, a little over 0.1 mm thick, made of cellulose acetate, cellulose nitrate, polycarbonate, polyvinylidene fluoride, or other synthetic materials. Although a wide variety of pore sizes are available, membranes with pores about 0.2 _m in diameter are used to remove most vegetative cells, but not viruses, from solutions ranging in volume from 1 ml to many liters. The membranes are held in special holders (figure) and often preceded by depth filters made of glass fibers to remove larger particles that might clog the membrane filter. The solution is pulled or forced through the filter with a vacuum or with pressure from a syringe, peristaltic pump, or nitrogen gas bottle, and collected in previously sterilized containers. Membrane filters remove microorganisms by screening them out much as a sieve separates large sand particles from small ones. These filters are used to sterilize pharmaceuticals, ophthalmic solutions, culture media, oils, antibiotics, and other heat-sensitive solutions.

Membrane Filter Sterilization. A membrane filter outfit for sterilizing medium volumes of solution. (a) Cross section of the membrane filtering unit. Several membranes are used to increase capacity. (b) A complete filtering setup. The solution to be sterilized is kept in the Erlenmeyer flask, 1, and forced through the filter by a peristaltic pump, 2. The solution is sterilized by flowing through a membrane filter unit, 3, and into a sterile container. A wide variety of other kinds of filtering outfits are also available.

Air also can be sterilized by filtration. Two common examples are surgical masks and cotton plugs on culture vessels that let air in but keep microorganisms out. Laminar flow biological safety cabinets employing high-efficiency particulate air (HEPA) filters, which remove 99.97% of 0.3 _m particles, are one of the most important air filtration systems. Laminar flow biological safety cabinets force air through HEPA filters, then project a vertical curtain of sterile air across the cabinet opening. This protects a worker from microorganisms being handled within the cabinet and prevents contamination of the room (figure). A person uses these cabinets when working with dangerous agents such as Mycobacterium tuberculosis, tumor viruses, and recombinant DNA. They are also employed in research labs and industries, such as the pharmaceutical industry, when a sterile working surface is needed for conducting assays, preparing media, examining tissue cultures, and the like.

A Laminar Flow Biological Safety Cabinet. A schematic diagram showing the airflow pattern.Describe each of the following agents in terms of its chemical nature, mechanism of action, mode of application, common uses and effectiveness, and advantages and disadvantages: phenolics, alcohols, halogens (iodine and chlorine), heavy metals, quaternary ammonium compounds, aldehydes, and ethylene oxide.Phenolics

Phenol was the first widely used antiseptic and disinfectant. In 1867 Joseph Lister employed it to reduce the risk of infection during operations. Today phenol and phenolics (phenol derivatives) such as cresols, xylenols, and orthophenylphenol are used as disinfectants in laboratories and hospitals. The commercial disinfectant Lysol is made of a mixture of phenolics. Phenolics act by denaturing proteins and disrupting cell membranes. They have some real advantages as disinfectants: phenolics are tuberculocidal, effective in the presence of organic material, and remain active on surfaces long after application. However, they do have a disagreeable odor and can cause skin irritation.

Hexachlorophene has been one of the most popular antiseptics because it persists on the skin once applied and reduces skin bacteria for long periods. However, it can cause brain damage and is now used in hospital nurseries only in response to a staphylococcal outbreak.3. Write down principle & application of TLC?

Thin Layer Chromatography - TLC

TLC is a simple, quick, and inexpensive procedure that gives the chemist a quick answer as to how many components are in a mixture. TLC is also used to support the identity of a compound in a mixture when the Rf of a compound is compared with the Rf of a known compound (preferrably both run on the same TLC plate).

A TLC plate is a sheet of glass, metal, or plastic which is coated with a thin layer of a solid adsorbent (usually silica or alumina). A small amount of the mixture to be analyzed is spotted near the bottom of this plate. The TLC plate is then placed in a shallow pool of a solvent in a developing chamber so that only the very bottom of the plate is in the liquid. This liquid, or the eluent, is the mobile phase, and it slowly rises up the TLC plate by capillary action.

As the solvent moves past the spot that was applied, an equilibrium is established for each component of the mixture between the molecules of that component which are adsorbed on the solid and the molecules which are in solution. In principle, the components will differ in solubility and in the strength of their adsorption to the adsorbent and some components will be carried farther up the plate than others. When the solvent has reached the top of the plate, the plate is removed from the developing chamber, dried, and the separated components of the mixture are visualized. If the compounds are colored, visualization is straightforward. Usually the compounds are not colored, so a UV lamp is used to visualize the plates. (The plate itself contains a fluor which fluoresces everywhere except where an organic compound is on the plate.)

The procedure for TLC, explained in words in the above paragraphs, is illustrated with photographs on the TLC Procedure page.

TLC Adsorbent

In the teaching labs at CU Boulder, we use silica gel plates (SiO2) almost exclusively. (Alumina (Al2O3) can also be used as a TLC adsorbent.) The plates are aluminum-backed and you can cut them to size with scissors. Our plates are purchased ready-made from EM Sciences or from Scientific Adsorbents. The adsorbent is impregnated with a fluor, zinc sulfide. The fluor enables most organic compounds to be visualized when the plate is held under a UV lamp. In some circumstances, other visualization methods are used, such as charring or staining.

TLC Solvents or Solvent Systems

Choosing a solvent is covered on the Chromatography Overview page. The charts at the bottom of that page are particularly useful.

Interactions of the Compound and the Adsorbent

The strength with which an organic compound binds to an adsorbent depends on the strength of the following types of interactions: ion-dipole, dipole-dipole, hydrogen bonding, dipole induced dipole, and van der Waals forces. With silica gel, the dominant interactive forces between the adsorbent and the materials to be separated are of the dipole-dipole type. Highly polar molecules interact fairly strongly with the polar Si—O bonds of these adsorbents and will tend to stick or adsorb onto the fine particles of the adsorbent while weakly polar molecules are held less tightly. Weakly polar molecules thus generally tend to move through the adsorbent more rapidly than the polar species. Roughly, the compounds follow the elution order given on the Chromatography Overview page.

The Rf value

Rf is the retention factor, or how far up a plate the compound travels. See the Rf page for more details:

Visualizing the Spots

If the compounds are colored, they are easy to see with the naked eye. If not, a UV lamp is used .

Troubleshooting TLC

All of the above (including the procedure page) might sound like TLC is quite an easy procedure. But what about the first time you run a TLC, and see spots everywhere and blurred, streaked spots? As with any technique, with practice you get better. One thing you have to be careful Examples of common problems encountered in TLC:

The compound runs as a streak rather than a spot

The sample was overloaded. Run the TLC again after diluting your sample. Or, your sample might just contain many components, creating many spots which run together and appear as a streak. Perhaps, the experiment did not go as well as expected.

The sample runs as a smear or a upward crescent.

Compounds which possess strongly acidic or basic groups (amines or carboxylic acids) sometimes show up on a TLC plate with this behavior. Add a few drops of ammonium hydroxide (amines) or acetic acid (carboxylic acids) to the eluting solvent to obtain clearer plates.

The sample runs as a downward crescent.

Likely, the adsorbent was disturbed during the spotting, causing the crescent shape.

The plate solvent front runs crookedly.

Either the adsorbent has flaked off the sides of the plate or the sides of the plate are touching the sides of the container (or the paper used to saturate the container) as the plate develops. Crookedly run plates make it harder to measure Rf values accurately.

Many, random spots are seen on the plate.

Make sure that you do not accidentally drop any organic compound on the plate. If get a TLC plate and leave it laying on your workbench as you do the experiment, you might drop or splash an organic compound on the plate.

No spots are seen on the plate.

You might not have spotted enough compound, perhaps because the solution of the compound is too dilute. Try concentrating the solution, or, spot it several times in one place, allowing the solvent to dry between applications. Some compounds do not show up under UV light; try another method of visualizing the plate. Or, perhaps you do not have any compound because your experiment did not go as well as planned.

If the solvent level in the developing jar is deeper than the origin (spotting line) of the TLC plate, the solvent will dissolve the compounds into the solvent reservoir instead of allowing them to move up the plate by capillary action. Thus, you will not see spots after the plate is developed.

4.Write down the principle & application of Column chromatography?

Column chromatography in chemistry is a method used to purify individual chemical compounds from mixtures of compounds. It is often used for preparative applications on scales from micrograms up to kilograms.

The classical preparative chromatography column is a glass tube with a diameter from 5 to 50 mm and a height of 50 cm to 1 m with a tap at the bottom. A slurry is prepared of the eluent with the stationary phase powder and then carefully poured into the column. Care must be taken to avoid air bubbles. A solution of the organic material is pipetted on top of the stationary phase. This layer is usually topped with a small layer of sand or with cotton or glass wool to protect the shape of the organic layer from the velocity of newly added eluant. Eluant is slowly passed through the column to advance the organic material. Often a spherical eluent reservoir or an eluent-filled and stoppered separating funnel is put on top of the column.

The individual components are retained by the stationary phase differently and separate from each other while they are running at different speeds through the column with the eluant. At the end of the column they elute one at a time. During the entire chromatography process the eluant is collected in a series of fractions. The composition of the eluant flow can be monitored and each fraction is analyzed for dissolved compounds, e.g. by analytical chromatography, UV absorption, or fluorescence. Colored compounds (or fluorescent compounds with the aid of an UV lamp) can be seen through the glass wall as moving bands.

Stationary phase (adsorbent)

The stationary phase or adsorbent in column chromatography is a solid. The most common stationary phase for column chromatography is silica gel, followed by alumina. Cellulose powder has often been used in the past. Also possible are ion exchange chromatography, reversed-phase chromatography (RP), affinity chromatography or expanded bed adsorption (EBA). The stationary phases are usually finely ground powders or gels and/or are microporous for an increased surface, though in EBA a fluidized bed is used.

Mobile phase (eluent)

The mobile phase or eluent is either a pure solvent or a mixture of different solvents. It is chosen so that the retention factor value of the compound of interest is roughly around 0.75 in order to minimize the time and the amount of eluent to run the chromatography. The eluent has also been chosen so that the different compounds can be separated effectively. The eluent is optimized in small scale pretests, often using thin layer chromatography (TLC) with the same stationary phase.

A faster flow rate of the eluent minimizes the time required to run a column and thereby minimizes diffusion, resulting in a better separation, see Van Deemter's equation. A simple laboratory column runs by gravity flow. The flow rate of such a column can be increased by extending the fresh eluent filled column above the top of the stationary phase or decreased by the tap controls. Better flow rates can be achieved by using a pump or by using compressed gas (e.g. air, nitrogen, or argon) to push the solvent through the column (flash column chromatography).[1]

A spreadsheet that assists in the successful development of flash columns has been developed. The spreadsheet estimates the retention volume and band volume of analytes, the fraction numbers expected to contain each analyte, and the resolution between adjacent peaks. This information allows users to select optimal parameters for preparative-scale separations before the flash column itself is attempted.[2]

Automated Systems

Column chromatography is an extremely time consuming stage in any lab and can quickly become the bottle neck for any process lab. Therefore, several manufactures have developed automated flash chromatography systems (typically referred to as LPLC, low pressure liquid chromatography, around 50-75 psi) that minimize human involvement in the purification process. Automated systems will include components normally found on more expensive HPLC systems such as a gradient pump, sample injection ports, a UV detector and a fraction collector to collect the eluent. Typically these automated systems can separate samples from a few milligrams up to an industrial kg scale and offer a much cheaper and quicker solution to doing multiple injections on prep-HPLC systems.

The resolution (or the ability to separate a mixture) on an LPLC system will always be lower compared to HPLC, as the packing material in an HPLC column can be much smaller, typically only 5 micrometre thus increasing stationary phase surface area, increasing surface interactions and giving better separation. However, the use of this small packing media causes the high back pressure and is why it is termed high pressure liquid chromatography. The LPLC columns are typically packed with silica of around 50 micrometres, thus reducing back pressure and resolution, but it also removes the need for expensive high pressure pumps. Manufactures are now starting to move into higher pressure flash chromatography systems and have termed these as medium pressure liquid chromatography (MPLC) systems which operate above 150 psi.

The software controlling an automated system will coordinate the components, allow a user to only collect the factions that contain their target compound (assuming they are detectable on the systems detector) and help the user to find the resulting purified material within the fraction collector. The software will also save the resulting chromatograph from the process for archival and/or later recall purposes.

A representative example of column chromatography as part of an undergraduate laboratory exercise is the separation of three components (out of 28) in the oil of spearmint: carvone, limonene and dehydrocarveol [3]. A microscale setup consisting of a

Pasteur pipette as column with silica gel stationary phase can suffice. The starting eluent is hexane and solvent polarity is increased during the process by adding ethyl acetate. Dr. bhakti

Column Chromatogram Resolution Calculation

Typically, column chromatography is set up with peristaltic pumps flowing buffers and the solution sample through the top of the column. The solutions and buffers pass through the column where a fraction collector at the end of the column setup collects the eluted samples from the it. Prior to the fraction collection, the samples that are eluted from the column pass through a detector such as a spectrophotometer or mass spectrometer so that the concentration of the separated samples in the sample solution mixture can be determined.

For example, if you were to separate two different proteins with different binding capacities to the column from a solution sample, a good type of detector would be a spectrophotometer using a wavelength of 280 nm. The higher the concentration of protein that passes through the eluted solution through the column, the higher the absorbance of that wavelength.Because the column chromatography has a constant flow of eluted solution passing through the detector at varying concentrations, the detector must plot the concentration of the eluted sample over a course of time. This plot of sample concentration versus time is called a chromatogram.

The ultimate goal of chromatography is to separate different components from a solution mixture. The resolution expresses the extent of separation between the components from the mixture. The higher the resolution of the chromatogram, the better the extent of separation of the samples the column gives. This data is a good way of determining the column’s separation properties of that particular sample. The resolution can be calculated from the chromatogram.The separate curves in the diagram represent different sample elution concentration profiles over time based on their affinity to the column resin. To calculate resolution, the retention time and curve width are required.

5.Explain in detail about UV &Visible spectrophotometer?

Ultraviolet-visible spectroscopy or ultraviolet-visible spectrophotometry (UV-Vis or UV/Vis) involves the spectroscopy of photons in the UV-visible region. It uses light in the visible and adjacent near ultraviolet (UV) and near infrared (NIR) ranges. In this region of the electromagnetic spectrum, molecules undergo electronic transitions. This technique is complementary to fluorescence spectroscopy, in that fluorescence deals with transitions from the excited state to the ground state, while absorption measures transitions from the ground state to the excited state.[1]

Ultraviolet-visible spectrum

An ultraviolet-visible spectrum is essentially a graph of light absorbance versus wavelength in a range of ultraviolet or visible regions. Such a spectrum can often be produced directly by a more sophisticated spectrophotometer, or the data can be collected one wavelength at a time by simpler instruments. Wavelength is often represented by the symbol λ. Similarly, for a given substance, a standard graph of the extinction coefficient (ε) vs. wavelength (λ) may be made or used if one is already available. Such a standard graph would be effectively "concentration-corrected" and thus independent of concentration.

The Woodward-Fieser rules are a set of empirical observations which can be used to predict λmax, the wavelength of the most intense UV/Vis absorption, for conjugated organic compounds such as dienes and ketones.

The wavelengths of absorption peaks can be correlated with the types of bonds in a given molecule and are valuable in determining the functional groups within a molecule. UV/Vis absorption is not, however, a specific test for any given compound. The nature of the solvent, the pH of the solution, temperature, high electrolyte concentrations, and the presence of interfering substances can influence the absorption spectra of compounds, as can variations in slit width (effective bandwidth) in the spectrophotometer.

Applications

UV/Vis spectroscopy is routinely used in the quantitative determination of solutions of transition metal ions and highly conjugated organic compounds.

Solutions of transition metal ions can be coloured (i.e., absorb visible light) because d electrons within the metal atoms can be excited from one electronic state to another. The colour of metal ion solutions is strongly affected by the presence of other species, such as certain anions or ligands. For instance, the colour of a dilute solution of copper sulfate is a very light blue; adding ammonia intensifies the colour and changes the wavelength of maximum absorption (λmax).

Organic compounds , especially those with a high degree of conjugation, also absorb light in the UV or visible regions of the electromagnetic spectrum. The solvents for these determinations are often water for water soluble compounds, or ethanol for organic-soluble compounds. (Organic solvents may have significant UV absorption; not all solvents are suitable for use in UV spectroscopy. Ethanol absorbs very weakly at most wavelengths.) Solvent polarity and pH can effect the absorption spectrum of an organic compound. Tyrosine, for example, increases in absorption maxima and molar extinction coefficient when pH increases from 6 to 13 or when solvent polarity decreases.

While charge transfer complexes also give rise to colours, the colours are often too intense to be used for quantitative measurement.

The Beer-Lambert law states that the absorbance of a solution is directly proportional to the solution's concentration. Thus UV/VIS spectroscopy can be used to determine the concentration of a solution. It is necessary to know how quickly the absorbance changes with concentration.

This can be taken from references (tables of molar extinction coefficients), or more accurately, determined from a calibration curve.

A UV/Vis spectrophotometer may be used as a detector for HPLC. The presence of an analyte gives a response which can be assumed to be proportional to the concentration. For accurate results, the instrument's response to the analyte in the unknown should be compared with the response to a standard; this is very similar to the use of calibration curves. The response (e.g., peak height) for a particular concentration is known as the response factor.

Beer-Lambert lawMain article: Beer-Lambert law

The method is most often used in a quantitative way to determine concentrations of an absorbing species in solution, using the Beer-Lambert law:

−,

where A is the measured absorbance, I0 is the intensity of the incident light at a given wavelength, I is the transmitted intensity, L the pathlength through the sample, and c the concentration of the absorbing species. For each species and wavelength, ε is a constant known as the molar absorptivity or extinction coefficient. This constant is a fundamental molecular property in a given solvent, at a particular temperature and pressure, and has units of 1 / M * cm or often AU / M * cm.

The absorbance and extinction ε are sometimes defined in terms of the natural logarithm instead of the base-10 logarithm.

The Beer-Lambert Law is useful for characterizing many compounds but does not hold as a universal relationship for the concentration and absorption of all substances. A 2nd order polynomial relationship between absorption and concentration is sometimes encountered for very large, complex molecules such as organic dyes (Xylenol Orange or Neutral Red, for example).

Practical Considerations

To actually make a valid measurement you must understand and be aware of the limitations of the particular instrument being used. This is especially important when making measurements using simple (and therefore relatively inexpensive) instruments, where a user is more likely to encounter an instrumental limitation, or when making measurements of materials that have not been well characterized yet.

The molar extinction coefficient, ε, is a function of the wavelength (that is, the color) of the light used. For the Beer-Lambert relation above to hold in a particular case, the light must be sufficiently monochromatic that the exctinction coefficient used is well defined.

For instance, the spectral bandwidth of the instrument (as FWHM), the portion of the spectrum selected for the measurement, must be much smaller than the width of the absorbance curve of

the sample, so that the exctintion coefficient does not change significantly over the band. Some instruments allow selection of bandwidth. (The tradeoff is that reducing the bandwidth reduces the energy passed to the detector and will require a longer measurement time to achieve the same signal to noise ratio.)

In liquids, the extinction coefficient usually changes slowly with wavelength. A peak of the absorbance curve (a wavelength where the absorbance reaches a maximum) is where the rate of change in absorbance with wavelength is smallest. Measurements are usually made at a peak in order to minimize errors produced by errors in wavelength in the instrument, that is errors due to having an different extinction coefficient than assumed.

Another important factor is the purity of the light used. The most important factor affecting this is the stray light level of the monochromator. The detector used is broadband, it responds to all the light that reaches it. If a significant amount of the light passed through the sample contains wavelengths that have much lower extinction coefficients than the nominal one, the instrument will report an incorrectly low absorbance. Any instrument will reach a point where an increase in sample concentration will not result in an increase in the reported absorbance, because the detector is simply responding to the stray light. In practice the concentration of the sample must be adjusted to place the unknown absorbance within a range that is valid for the instrument. Sometimes an empirical calibration function is developed, using known concentrations of the sample, to allow measurements into the region where the instrument is becoming non-linear.

Ultraviolet-visible spectrophotometerSee also: Spectrophotometry

The instrument used in ultraviolet-visible spectroscopy is called a UV/vis spectrophotometer. It measures the intensity of light passing through a sample (I), and compares it to the intensity of light before it passes through the sample (Io). The ratio I / Io is called the transmittance, and is usually expressed as a percentage (%T). The absorbance, A, is based on the transmittance:

A = − log(%T)

The basic parts of a spectrophotometer are a light source, a holder for the sample, a diffraction grating or monochromator to separate the different wavelengths of light, and a detector. The radiation source is often a Tungsten filament (300-2500 nm), a deuterium arc lamp which is continuous over the ultraviolet region (190-400 nm), and more recently light emitting diodes (LED) and Xenon Arc Lamps[2] for the visible wavelengths. The detector is typically a photodiode or a CCD. Photodiodes are used with monochromators, which filter the light so that only light of a single wavelength reaches the detector. Diffraction gratings are used with CCDs, which collects light of different wavelengths on different pixels.

Diagram of a single-beam UV/vis spectrophotometer.

A spectrophotometer can be either single beam or double beam. In a single beam instrument (such as the Spectronic 20), all of the light passes through the sample cell. Io must be measured

by removing the sample. This was the earliest design, but is still in common use in both teaching and industrial labs.

In a double-beam instrument, the light is split into two beams before it reaches the sample. One beam is used as the reference; the other beam passes through the sample. Some double-beam instruments have two detectors (photodiodes), and the sample and reference beam are measured at the same time. In other instruments, the two beams pass through a beam chopper, which blocks one beam at a time. The detector alternates between measuring the sample beam and the reference beam.

Samples for UV/Vis spectrophotometry are most often liquids, although the absorbance of gases and even of solids can also be measured. Samples are typically placed in a transparent cell, known as a cuvette. Cuvettes are typically rectangular in shape, commonly with an internal width of 1 cm. (This width becomes the path length, L, in the Beer-Lambert law.) Test tubes can also be used as cuvettes in some instruments. The type of sample container used must allow radiation to pass over the spectral region of interest. The most widely applicable cuvettes are made of high quality fused silica or quartz glass because these are transparent throughout the UV, visible and near infrared regions. Glass and plastic cuvettes are also common, although glass and most plastics absorb in the UV, which limits their usefulness to visible wavelengths.[3]

The wavelengths of absorption peaks can be correlated with the types of bonds in a given molecule and are valuable in determining the functional groups within a molecule. UV/Vis absorption is not, however, a specific test for any given compound. The nature of the solvent, the pH of the solution, temperature, high electrolyte concentrations, and the presence of interfering substances can influence the absorption spectra of compounds, as can variations in slit width (effective bandwidth) in the spectrophotometer.

Optical System Diagram

The UV-Visible spectrophotometer uses two light sources, a deuterium (D2) lamp for ultraviolet light and a tungsten (W) lamp for visible light. After bouncing off a mirror (mirror 1), the light beam passes through a slit and hits a diffraction grating. The grating can be rotated allowing for a specific wavelength to be selected. At any specific orientation of the grating, only monochromatic (single wavelength) successfully passes through a slit. A filter is used to remove unwanted higher orders of diffraction. (Recall the experiment you did last semester on Atomic Spectra) The light beam hits a second mirror before it gets split by a half mirror (half of the light is reflected, the other half passes through). One of the beams is allowed to pass through a reference cuvette (which contains the solvent only), the other passes through the sample cuvette. The intensities of the light beams are then measured at the end.

Beer-Lambert Law

The change in intensity of light (dI) after passing through a sample should be proportional to the following:

(a) path length (b), the longer the path, more photons should be absorbed

(b) concentration (c) of sample, more molecules absorbing means more photons absorbed

(c) intensity of the incident light (I), more photons mean more opportunity for a molecule to see a photon

Thus,

dI is proportional to bcI or

dI/I = -kbc (where k is a proportionality constant, the negative sign is shown because this is a decrease in intensity of the light, this makes b, c and I always positive.

Integration of the above equation leads to Beer-Lambert's Law

6.Explain in detail about Ion exchange chromatography?

Ion-exchange chromatography (or ion chromatography) is a process that allows the separation of ions and polar molecules based on the charge properties of the molecules. It can be used for almost any kind of charged molecule including large proteins, small nucleotides and amino acids. The solution to be injected is usually called a sample, and the individually separated components are called analytes. It is often used in protein purification, water analysis, and quality control

History

Ion methods have been in use since 1850, when H. Thompson and J. T. Way, researchers in England, treated various clays with ammonium sulfate or carbonate in solution to extract the ammonia and release calcium. In 1927, the first zeolite mineral column was used to remove interfering calcium and magnesium ions from solution to determine the sulfate content of water. The modern version of IEC was developed during the wartime Manhattan Project. A technique was required to separate and concentrate the radioactive elements needed to make the atom bomb. Researchers chose adsorbents that would latch onto charged transuranium elements, which could then be differentially eluted. Ultimately, once declassified, these techniques would use new IE resins to develop the systems that are often used today for specific purification of biologicals and inorganics. In the early 1970s, ion chromatography was developed by Hamish Small and co-workers at Dow Chemical Company as a novel method of IEC usable in automated analysis. IC uses weaker ionic resins for its stationary phase and an additional neutralizing stripper, or suppressor, column to remove background eluent ions. It is a powerful technique for

determining low concentrations of ions and is especially useful in environmental and water quality studies, among other applications.

The Dow Chemical Company technology was acquired by Durrum Instrument Corp. (maker of the Durrum D-500), which later formed a separate business unit for its new IC products, naming it Dionex (Dow Ion Exchange). Dionex Corporation was incorporated in Sunnyvale, California in 1980, and, led by A. Blaine Bowman, purchased the Dionex assets.

Principle

Ion Chromatogram

Ion exchange chromatography retains analyte molecules based on coulombic (ionic) interactions. The stationary phase surface displays ionic functional groups (R-X) that interact with analyte ions of opposite charge. This type of chromatography is further subdivided into cation exchange chromatography and anion exchange chromatography. The ionic compound consisting of the cationic species M+ and the anionic species B- can be retained by the stationary phase.

Cation exchange chromatography retains positively charged cations because the stationary phase displays a negatively charged functional group:

Anion exchange chromatography retains anions using positively charged functional group:

Note that the ion strength of either C+ or A- in the mobile phase can be adjusted to shift the equilibrium position and thus retention time.

The ion chromatogram shows a typical chromatogram obtained with an anion exchange column.

Typical technique

Another ion chromatography workstation

A sample is introduced, either manually or with an autosampler, into a sample loop of known volume. A buffered aqueous solution known as the mobile phase carries the sample from the loop onto a column that contains some form of stationary phase material. This is typically a resin or gel matrix consisting of agarose or cellulose beads with covalently bonded charged functional groups. The target analytes (anions or cations) are retained on the stationary phase but can be eluted by increasing the concentration of a similarly charged species that will displace the analyte ions from the stationary phase. For example, in cation exchange chromatography, the positively charged analyte could be displaced by the addition of positively charged sodium ions. The analytes of interest must then be detected by some means, typically by conductivity or UV/Visible light absorbance.

In order to control an IC system, a chromatography data system (CDS) is usually needed. In addition to IC systems, some of these CDSs can also control gas chromatography (GC) and HPLC systems.

Separating proteins

Preparative-scale ion exchange column used for protein purification.

Proteins have numerous functional groups that can have both positive and negative charges. Ion exchange chromatography separates proteins according to their net charge, which is dependent on the composition of the mobile phase. By adjusting the pH or the ionic concentration of the mobile phase, various protein molecules can be separated. For example, if a protein has a net positive charge at pH 7, then it will bind to a column of negatively-charged beads, whereas a negatively charged protein would not. By changing the pH so that the net charge on the protein is negative, it too will be eluted.

Elution by changing the ionic strength of the mobile phase is a more subtle effect - it works as ions from the mobile phase will interact with the immobilized ions in preference over those on the stationary phase. This "shields" the stationary phase from the protein, (and vice versa) and allows the protein to elute

7. Explain in detail about Electrophoresis?

Electrophoresis is the best-known electrokinetic phenomenon. It was discovered by Reuss in 1807.[1] He observed that clay particles dispersed in water migrate under influence of an applied electric field. There are detailed descriptions of Electrophoresis in many books on Colloid and Interface Science.[2][3][4][5][6][7] There is an IUPAC Technical Report[8] prepared by a group of well known experts on the electrokinetic phenomena. Generally, electrophoresis is the motion of dispersed particles relative to a fluid under the influence of an electric field that is space uniform. Alternatively, similar motion in a space non-uniform electric field is called dielectrophoresis.

Electrophoresis occurs because particles dispersed in a fluid almost always carry an electric surface charge. An electric field exerts electrostatic Coulomb force on the particles through these charges. Recent molecular dynamics simulations, though, suggest that surface charge is not always necessary for electrophoresis and that even neutral particles can show electrophoresis due to the specific molecular structure of waterer at the interface.[9]

The electrostatic Coulomb force exerted on a surface charge is reduced by an opposing force which is electrostatic as well. According to double layer theory, all surface charges in fluids are screened by a diffuse layer. This diffuse layer has the same absolute charge value, but with opposite sign from the surface charge. The electric field induces force on the diffuse layer, as well as on the surface charge. The total value of this force equals to the first mentioned force, but it is oppositely directed. However, only part of this force is applied to the particle. It is actually applied to the ions in the diffuse layer. These ions are at some distance from the particle surface.

They transfer part of this electrostatic force to the particle surface through viscous stress. This part of the force that is applied to the particle body is called electrophoretic retardation force.

There is one more electric force, which is associated with deviation of the double layer from spherical symmetry and surface conductivity due to the excess ions in the diffuse layer. This force is called the electrophoretic relaxation force.

All these forces are balanced with hydrodynamic friction, which affects all bodies moving in viscous fluids with low Reynolds number. The speed of this motion v is proportional to the electric field strength E if the field is not too strong. Using this assumption makes possible the introduction of electrophoretic mobility μe as coefficient of proportionality between particle speed and electric field strength:

Multiple theories were developed during 20th century for calculating this parameter. Ref. 2 provides an overview.

Theory

The most known and widely used theory of electrophoresis was developed by Smoluchowski in 1903 [10]

,

where ε is the dielectric constant of the dispersion medium, ε0 is the permittivity of free space (C² N-1 m-2), η is dynamic viscosity of the dispersion medium (Pa s), and ζ is zeta potential (i.e., the electrokinetic potential of the slipping plane in the double layer).

Smoluchowski theory is very powerful because it works for dispersed particles of any shape and any concentration, when it is valid. Unfortunately, it has limitations of its validity. It follows, for

instance, from the fact that it does not include Debye length κ-1. However, Debye length must be important for electrophoresis, as follows immediately from the Figure on the right. Increasing thickness of the DL leads to removing point of retardation force further from the particle surface. The thicker DL, the smaller retardation force must be.

Detailed theoretical analysis proved that Smoluchowski theory is valid only for sufficiently thin DL, when Debye length is much smaller than particle radius a:

κa > > 1

This model of "thin Double Layer" offers tremendous simplifications not only for electrophoresis theory but for many other electrokinetic theories. This model is valid for most aqueous systems because the Debye length is only a few nanometers there. It breaks only for nano-colloids in solution with ionic strength close to water

Smoluchowski theory also neglects contribution of surface conductivity. This is expressed in modern theory as condition of small Dukhin number

Du < < 1

Creation of electrophoretic theory with wider range of validity was a purpose of many studies during 20th century.

One of the most known considers an opposite asymptotic case when Debye length is larger than particle radius:

κa < 1

It is called the "thick Double Layer" model. Corresponding electrophoretic theory was created by Huckel in 1924 [11]. It yields the following equation for electrophoretic mobility:

,

This model can be useful for some nano-colloids and non-polar fluids, where Debye length is much larger.

There are several analytical theories that incorporate surface conductivity and eliminate restriction of the small Dukhin number. Early pioneering work in that direction dates back to Overbeek [12] and Booth [13].

Modern, rigorous theories that are valid for any Zeta potential and often any κa, stem mostly from the Ukrainian (Dukhin, Shilov and others) and Australian (O'Brien, White, Hunter and others) Schools.

Historically the first one was Dukhin-Semenikhin theory [14]. Similar theory was created 10 years later by O'Brien and Hunter [15]. Assuming thin Double Layer, these theories would yield results that are very close to the numerical solution provided by O'Brien and White [

8.Write a note on Gas liquid chromatography?

Gas-liquid chromatography (GLC), or simply gas chromatography (GC), is a type of chromatography in which the mobile phase is a carrier gas, usually an inert gas such as helium or an unreactive gas such as nitrogen, and the stationary phase is a microscopic layer of liquid or polymer on an inert solid support, inside glass or metal tubing, called a column. The instrument used to perform gas chromatographic separations is called a gas chromatograph (also: aerograph, gas separator).

Gas Chromatography is different from other forms of chromatography (HPLC, TLC, etc.) because the solutions travel through the column in a gas state. The interactions of these gaseous analytes with the walls of the column (coated by different stationary phases) causes different compounds to elute at different times called retention time. The comparison of these retention times is the analytical power of GC. This makes it very similar to high performance liquid chromatography.

History

Chromatography dates to 1903 in the work of the Russian scientist, Mikhail Semenovich Tswett. German graduate student Fritz Prior developed solid state gas chromatography in 1947. Archer John Porter Martin, who was awarded the Nobel Prize for his work in developing liquid-liquid (1941) and paper (1944) chromatography, laid the foundation for the development of gas chromatography and later produced liquid-gas chromatography (1950).

GC analysis

A gas chromatograph is a chemical analysis instrument for separating chemicals in a complex sample. A gas chromatograph uses a flow-through narrow tube known as the column, through which different chemical constituents of a sample pass in a gas stream (carrier gas, mobile phase) at different rates depending on their various chemical and physical properties and their interaction with a specific column filling, called the stationary phase. As the chemicals exit the end of the column, they are detected and identified electronically. The function of the stationary phase in the column is to separate different components, causing each one to exit the column at a different time (retention time). Other parameters that can be used to alter the order or time of retention are the carrier gas flow rate, and the temperature.

In a GC analysis, a known volume of gaseous or liquid analyte is injected into the "entrance" (head) of the column, usually using a microsyringe (or, solid phase microextraction fibers, or a gas source switching system). As the carrier gas sweeps the analyte molecules through the column, this motion is inhibited by the adsorption of the analyte molecules either onto the

column walls or onto packing materials in the column. The rate at which the molecules progress along the column depends on the strength of adsorption, which in turn depends on the type of molecule and on the stationary phase materials. Since each type of molecule has a different rate of progression, the various components of the analyte mixture are separated as they progress along the column and reach the end of the column at different times (retention time). A detector is used to monitor the outlet stream from the column; thus, the time at which each component reaches the outlet and the amount of that component can be determined. Generally, substances are identified (qualitatively) by the order in which they emerge (elute) from the column and by the retention time of the analyte in the column.

Physical componentsDiagram of a gas chromatograph.

Autosamplers

The autosampler provides the means to introduce automatically a sample into the inlets. Manual insertion of the sample is possible but is no longer common. Automatic insertion provides better reproducibility and time-optimization.

Different kinds of autosamplers exist. Autosamplers can be classified in relation to sample capacity (auto-injectors VS autosamplers, where auto-injectors can work a small number of samples), to robotic technologies (XYZ robot VS rotating/SCARA-robot – the most common), or to analysis:

Liquid Static head-space by syringe technology Dynamic head-space by transfer-line technology SPME

Traditionally autosampler manufactures are different from GC manufactures and currently no GC manufacture offers a complete range of autosamplers. Historically, the countries most active in autosampler technology development are the United States, Italy, and Switzerland.

Inlets

The column inlet (or injector) provides the means to introduce a sample into a continuous flow of carrier gas. The inlet is a piece of hardware attached to the column head.

Common inlet types are:

S/SL (Split/Splitless) injector; a sample is introduced into a heated small chamber via a syringe through a septum - the heat facilitates volatilization of the sample and sample matrix. The carrier gas then either sweeps the entirety (splitless mode) or a portion (split mode) of the sample into the column. In split mode, a part of the sample/carrier gas mixture in the injection chamber is exhausted through the split vent.

On-column inlet; the sample is here introduced in its entirety without heat.

PTV injector; Temperature-programmed sample introduction was first described by Vogt in 1979. Originally Vogt developed the technique as a method for the introduction of large sample volumes (up to 250 µL) in capillary GC. Vogt introduced the sample into the liner at a controlled injection rate. The temperature of the liner was chosen slightly below the boiling point of the solvent. The low-boiling solvent was continuously evaporated and vented through the split line. Based on this technique, Poy developed the Programmed Temperature Vaporising injector; PTV. By introducing the sample at a low initial liner temperature many of the disadvantages of the classic hot injection techniques could be circumvented.

Gas source inlet or gas switching valve; gaseous samples in collection bottles are connected to what is most commonly a six-port switching valve. The carrier gas flow is not interrupted while a sample can be expanded into a previously evacuated sample loop. Upon switching, the contents of the sample loop are inserted into the carrier gas stream.

P/T (Purge-and-Trap) system; An inert gas is bubbled through an aqueous sample causing insoluble volatile chemicals to be purged from the matrix. The volatiles are 'trapped' on an absorbent column (known as a trap or concentrator) at ambient temperature. The trap is then heated and the volatiles are directed into the carrier gas stream. Samples requiring preconcentration or purification can be introduced via such a system, usually hooked up to the S/SL port.

SPME (solid phase microextraction) offers a convenient, low-cost alternative to P/T systems with the versatility of a syringe and simple use of the S/SL port.

Columns

Two types of columns are used in GC:

Packed columns are 1.5 - 10 m in length and have an internal diameter of 2 - 4 mm. The tubing is usually made of stainless steel or glass and contains a packing of finely divided, inert, solid support material (eg. diatomaceous earth) that is coated with a liquid or solid stationary phase. The nature of the coating material determines what type of materials will be most strongly adsorbed. Thus numerous columns are available that are designed to separate specific types of compounds.

Capillary columns have a very small internal diameter, on the order of a few tenths of millimeters, and lengths between 25-60 meters are common. The inner column walls are coated with the active materials (WCOT columns), some columns are quasi solid filled with many parallel micropores (PLOT columns). Most capillary columns are made of fused-silica with a polyimide outer coating. These columns are flexible, so a very long column can be wound into a small coil.

New developments are sought where stationary phase incompatibilities lead to geometric solutions of parallel columns within one column. Among these new developments are:

o Internally heated microFAST columns, where two columns, an internal heating wire and a temperature sensor are combined within a common column sheath (microFAST);

o Micropacked columns (1/16" OD) are column-in-column packed columns where the outer column space has a packing different from the inner column space, thus

providing the separation behaviour of two columns in one. They can easily fit to inlets and detectors of a capillary column instrument.

The temperature-dependence of molecular adsorption and of the rate of progression along the column necessitates a careful control of the column temperature to within a few tenths of a degree for precise work. Reducing the temperature produces the greatest level of separation, but can result in very long elution times. For some cases temperature is ramped either continuously or in steps to provide the desired separation. This is referred to as a temperature program. Electronic pressure control can also be used to modify flow rate during the analysis, aiding in faster run times while keeping acceptable levels of separation.

The choice of carrier gas (mobile phase) is important, with hydrogen being the most efficient and providing the best separation. However, helium has a larger range of flowrates that are comparable to hydrogen in efficiency, with the added advantage that helium is non-flammable, and works with a greater number of detectors. Therefore, helium is the most common carrier gas used.

Detectors

A number of detectors are used in gas chromatography. The most common are the flame ionization detector (FID) and the thermal conductivity detector (TCD). Both are sensitive to a wide range of components, and both work over a wide range of concentrations. While TCDs are essentially universal and can be used to detect any component other than the carrier gas (as long as their thermal conductivities are different from that of the carrier gas, at detector temperature), FIDs are sensitive primarily to hydrocarbons, and are more sensitive to them than TCD. However, an FID cannot detect water. Both detectors are also quite robust. Since TCD is non-destructive, it can be operated in-series before an FID (destructive), thus providing complementary detection of the same analytes.

Other detectors are sensitive only to specific types of substances, or work well only in narrower ranges of concentrations. They include:

discharge ionization detector (DID), which uses a high-voltage electric discharge to produce ions.

electron capture detector (ECD), which uses a radioactive Beta particle (electron) source to measure the degree of electron capture.

flame photometric detector (FPD) flame ionization detector (FID) Hall electrolytic conductivity detector (ElCD) helium ionization detector (HID) nitrogen phosphorus detector (NPD) mass selective detector (MSD) photo-ionization detector (PID) pulsed discharge ionization detector (PDD) thermal energy(conductivity) analyzer/detector (TEA/TCD)

Some gas chromatographs are connected to a mass spectrometer which acts as the detector. The combination is known as GC-MS. Some GC-MS are connected to an NMR spectrometer which acts as a back up detector. This combination is known as GC-MS-NMR. Some GC-MS-NMR are connected to an infrared spectrophotometer which acts as a back up detector. This combination is known as GC-MS-NMR-IR. It must, however, be stressed this is very rare as most analyses needed can be concluded via purely GC-MS.

Methods

The method is the collection of conditions in which the GC operates for a given analysis. Method development is the process of determining what conditions are adequate and/or ideal for the analysis required.

Conditions which can be varied to accommodate a required analysis include inlet temperature, detector temperature, column temperature and temperature program, carrier gas and carrier gas flow rates, the column's stationary phase, diameter and length, inlet type and flow rates, sample size and injection technique. Depending on the detector(s) (see below) installed on the GC, there may be a number of detector conditions that can also be varied. Some GCs also include valves which can change the route of sample and carrier flow. The timing of the opening and closing of these valves can be important to method development.

This image above shows the interior of a GeoStrata Technologies Eclipse Gas Chromatograph that runs continuously in three minute cycles. Two valves are used to switch the test gas into the sample loop. After filling the sample loop with test gas, the valves are switched again applying carrier gas pressure to the sample loop and forcing the sample through the Column for separation.

Carrier gas selection and flow rates

Typical carrier gases include helium, nitrogen, argon, hydrogen and air. Which gas to use is usually determined by the detector being used, for example, a DID requires helium as the carrier gas. When analyzing gas samples, however, the carrier is sometimes selected based on the sample's matrix, for example, when analyzing a mixture in argon, an argon carrier is preferred, because the argon in the sample does not show up on the chromatogram. Safety and availability

can also influence carrier selection, for example, hydrogen is flammable, and high-purity helium can be difficult to obtain in some areas of the world. (See: Helium--occurrence and production.)

The purity of the carrier gas is also frequently determined by the detector, though the level of sensitivity needed can also play a significant role. Typically, purities of 99.995% or higher are used. Trade names for typical purities include "Zero Grade," "Ultra-High Purity (UHP) Grade," "4.5 Grade" and "5.0 Grade."

The carrier gas flow rate affects the analysis in the same way that temperature does (see above). The higher the flow rate the faster the analysis, but the lower the separation between analytes. Selecting the flow rate is therefore the same compromise between the level of separation and length of analysis as selecting the column temperature.

With GCs made before the 1990s, carrier flow rate was controlled indirectly by controlling the carrier inlet pressure, or "column head pressure." The actual flow rate was measured at the outlet of the column or the detector with an electronic flow meter, or a bubble flow meter, and could be an involved, time consuming, and frustrating process. The pressure setting was not able to be varied during the run, and thus the flow was essentially constant during the analysis. The relation between flow rate and inlet pressure is calculated with Poiseuille's equation for compressible fluids.

Many modern GCs, however, electronically measure the flow rate, and electronically control the carrier gas pressure to set the flow rate. Consequently, carrier pressures and flow rates can be adjusted during the run, creating pressure/flow programs similar to temperature programs.

Inlet types and flow rates

The choice of inlet type and injection technique depends on if the sample is in liquid, gas, adsorbed, or solid form, and on whether a solvent matrix is present that has to be vaporized. Dissolved samples can be introduced directly onto the column via a COC injector, if the conditions are well known; if a solvent matrix has to be vaporized and partially removed, a S/SL injector is used (most common injection technique); gaseous samples (e.g., air cylinders) are usually injected using a gas switching valve system; adsorbed samples (e.g., on adsorbent tubes) are introduced using either an external (on-line or off-line) desorption apparatus such as a purge-and-trap system, or are desorbed in the S/SL injector (SPME applications).

Sample size and injection technique

Sample injection

The rule of ten in gas chromatography

The real chromatographic analysis starts with the introduction of the sample onto the column. The development of capillary gas chromatography resulted in many practical problems with the injection technique. The technique of on-column injection, often used with packed columns, is usually not possible with capillary columns. The injection system, in the capillary gas chromatograph, should fulfil the following two requirements:

1. The amount injected should not overload the column. 2. The width of the injected plug should be small compared to the spreading due to the

chromatographic process. Failure to comply with this requirement will reduce the separation capability of the column. As a general rule, the volume injected, V inj, and the volume of the detector cell, Vdet, should be about 1/10 of the volume occupied by the portion of sample containing the molecules of interest (analytes) when they exit the column.

Some general requirements, which a good injection technique should fulfill, are:

It should be possible to obtain the column’s optimum separation efficiency. It should allow accurate and reproducible injections of small amounts of representative

samples. It should induce no change in sample composition. It should not exhibit discrimination

based on differences in boiling point, polarity, concentration or thermal/catalytic stability. It should be applicable for trace analysis as well as for undiluted samples.

Column selection

Column temperature and temperature program

A gas chromatography oven, open to show a capillary column

The column(s) in a GC are contained in an oven, the temperature of which is precisely controlled electronically. (When discussing the "temperature of the column," an analyst is technically referring to the temperature of the column oven. The distinction, however, is not important and will not subsequently be made in this article.)

The rate at which a sample passes through the column is directly proportional to the temperature of the column. The higher the column temperature, the faster the sample moves through the column. However, the faster a sample moves through the column, the less it interacts with the stationary phase, and the less the analytes are separated.

In general, the column temperature is selected to compromise between the length of the analysis and the level of separation.

A method which holds the column at the same temperature for the entire analysis is called "isothermal." Most methods, however, increase the column temperature during the analysis, the initial temperature, rate of temperature increase (the temperature "ramp") and final temperature is called the "temperature program."

A temperature program allows analytes that elute early in the analysis to separate adequately, while shortening the time it takes for late-eluting analytes to pass through the column.

Data reduction and analysis

Qualitative analysis:

Generally chromatographic data is presented as a graph of detector response (y-axis) against retention time (x-axis), which is called a chromatogram. This provides a spectrum of peaks for a sample representing the analytes present in a sample eluting from the column at different times. Retention time can be used to identify analytes if the method conditions are constant. Also, the

pattern of peaks will be constant for a sample under constant conditions and can identify complex mixtures of analytes. In most modern applications however the GC is connected to a mass spectrometer or similar detector that is capable of identifying the analytes represented by the peaks.

Quantitive analysis:

The area under a peak is proportional to the amount of analyte present in the chromatogram. By calculating the area of the peak using the mathematical function of integration, the concentration of an analyte in the original sample can be determined. Concentration can be calculated using a calibration curve created by finding the response for a series of concentrations of analyte, or by determining the relative response factor of an analyte. The relative response factor is the expected ratio of an analyte to an internal standard (or external standard) and is calculated by finding the response of a known amount of analyte and a constant amount of internal standard (a chemical added to the sample at a constant concentration, with a distinct retention time to the analyte).

In most modern GC-MS systems, computer software is used to draw and integrate peaks, and match MS spectra to library spectra.

Application

In general, substances that vaporize below ca. 300 °C (and therefore are stable up to that temperature) can be measured quantitatively. The samples are also required to be salt-free; they should not contain ions. Very minute amounts of a substance can be measured, but it is often required that the sample must be measured in comparison to a sample containing the pure, suspected substance.

Various temperature programs can be used to make the readings more meaningful; for example to differentiate between substances that behave similarly during the GC process.

Professionals working with GC analyze the content of a chemical product, for example in assuring the quality of products in the chemical industry; or measuring toxic substances in soil, air or water. GC is very accurate if used properly and can measure picomoles of a substance in a 1 ml liquid sample, or parts-per-billion concentrations in gaseous samples.

In practical courses at colleges, students sometimes get acquainted to the GC by studying the contents of Lavender oil or measuring the ethylene that is secreted by Nicotiana benthamiana plants after artificially injuring their leaves. These GC analyses hydrocarbons (C2-C40+). In a typical experiment, a packed column is used to separate the light gases, which are then detected with a TCD. The hydrocarbons are separated using a capillary column and detected with an FID. A complication with light gas analyses that include H2 is that He, which is the most common and most sensitive inert carrier (sensitivity is proportional to molecular mass) has an almost identical thermal conductivity to hydrogen (it is the difference in thermal conductivity between two separate filaments in a Wheatstone Bridge type arrangement that shows when a component has been eluted). For this reason, dual TCD instruments are used with a separate channel for

hydrogen that uses nitrogen as a carrier are common. Argon is often used when analysing gas phase chemistry reactions such as F-T synthesis so that a single carrier gas can be used rather than 2 separate ones. The sensitivity is less but this is a tradeoff for simplicity in the gas supply.

GCs in popular culture

Movies, books and TV shows tend to misrepresent the capabilities of gas chromatography and the work done with these instruments.

In the U.S. TV show CSI, for example, GCs are used to rapidly identify unknown samples. "This is gasoline bought at a Chevron station in the past two weeks," the analyst will say fifteen minutes after receiving the sample.

In fact, a typical GC analysis takes much more time; sometimes a single sample must be run more than an hour according to the chosen program; and even more time is needed to "heat out" the column so it is free from the first sample and can be used for the next. Equally, several runs are needed to confirm the results of a study - a GC analysis of a single sample may simply yield a result per chance (see statistical significance).

Also, GC does not positively identify most samples; and not all substances in a sample will necessarily be detected. All a GC truly tells you is at which relative time a component eluted from the column and that the detector was sensitive to it. To make results meaningful, analysts need to know which components at which concentrations are to be expected; and even then a small amount of a substance can hide itself behind a substance having both a higher concentration and the same relative elution time. Last but not least it is often needed to check the results of the sample against a GC analysis of a reference sample containing only the suspected substance.

A GC-MS can remove much of this ambiguity, since the mass spectrometer will identify the component's molecular weight. But this still takes time and skill to do properly.

Similarly, most GC analyses are not push-button operations. You cannot simply drop a sample vial into an auto-sampler's tray, push a button and have a computer tell you everything you need to know about the sample. According to the substances one expects to find the operating program must be carefully chosen.

A push-button operation can exist for running similar samples repeatedly, such as in a chemical production environment or for comparing 20 samples from the same experiment to calculate the mean content of the same substance. However, for the kind of investigative work portrayed in books, movies and TV shows this is clearly not the case.

9.Write down principle & application of Autoclave?

Autoclave

Autoclave

A modern front-loading autoclave

Uses Sterilization

Inventor Charles Chamberland

Related items Waste autoclave

An autoclave is a pressurized device designed to heat aqueous solutions above their boiling point at normal atmospheric pressure to achieve sterilization. It was invented by Charles Chamberland in 1879.[1] The term autoclave is also used to describe an industrial machine in which elevated temperature and pressure are used in processing materials.

Introduction

Under ordinary circumstances (at standard pressure), liquid water cannot be heated above approximately 100 °C/212 °F (99.99 °C at 101.325 kPa, 99.62 °C at 100 kPa) in an open vessel (see here for special situations). Further heating results in boiling, which is the transition from liquid to gas, but does not raise the temperature of the liquid water. However, when water is heated in a pressurized vessel such as an autoclave, it is possible to heat liquid water to a much higher temperature. As the container is heated the pressure rises due to the constant volume of

the container (see the ideal gas law). The boiling point of the water is raised because the amount of energy needed to form steam against the higher pressure is increased.

Uses

Autoclaves are widely used in microbiology, medicine, sterilizing instruments for body piercing, veterinary science, dentistry, podiatry and metallurgy. The large carbon-fiber composite parts for the Boeing 787, such as wing and fuselage parts, are cured in large autoclaves.[2]

Air removal

When the goal of autoclaving is to achieve sterility, it is very important to ensure that all of the trapped air is removed. The reason for this is that 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.[citation needed] Autoclaves may achieve air removal by various means including:

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 - Some autoclaves remove air by using a series of steam pulses, in which the chamber is alternately pressurized and then depressurized to near atmospheric pressure.

Vacuum pumps - Some autoclaves use vacuum pumps 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.

Autoclaves in medicine

Stovetop autoclaves - the simplest of autoclaves

A medical autoclave is a device that uses steam to sterilize equipment and other objects. This means that all bacteria, viruses, fungi, and spores are inactivated. However, prions, like those associated with Creutzfeldt-Jakob disease, may not be destroyed by autoclaving at the typical 121 °C for 15 minutes or 134 °C for 3 minutes, but can be destroyed with a longer sterilization cycle of 134 °C for 18 minutes[citation needed]. Also, some recently-discovered organisms, such as Strain 121, can survive at temperatures above 121 °C.

Autoclaves are found in many medical settings and other places that need to ensure sterility of an object. Many procedures today use single-use items rather than sterilized, reusable items. This first happened with hypodermic needles, but today many surgical instruments (such as forceps, needle holders, and scalpel handles) are commonly single-use items rather than reusable. See waste autoclave.

Because damp heat is used, heat-labile products (such as some plastics) cannot be sterilized this way or they will melt. Some paper or other products that may be damaged by the steam must also be sterilized another way. In all autoclaves, items should always be separated to allow the steam to penetrate the load evenly.

Autoclaving is often used to sterilize medical waste prior to disposal in the standard municipal solid waste stream. This application has grown as an alternative to incineration due to environmental and health concerns raised by combustion byproducts from incinerators, especially from the small units which were commonly operated at individual hospitals. Incineration or a similar thermal oxidation process is still generally mandated for pathological waste and other very toxic and/or infectious medical wastes.

Autoclave quality assurance

Multiple large autoclaves are used for processing substantial quantities of laboratory equipment prior to reuse, and infectious material prior to disposal. The machine in the middle is a washing machine, the machine to the right is the Autoclave

Sterilization bags often have a "sterilization indicator mark" that typically darkens when sterilization temperatures have been reached. Comparing the mark on an unprocessed bag (L) to a bag that has been properly cycled (R) will show an obvious visual difference.

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 autoclaved sufficiently. Biological indicators include attest devices. These contain spores of a heat-resistant bacterium, Geobacillus stearothermophilus. If the autoclave does not reach the right temperature, the spores will germinate, and their metabolism will change the color of a pH-sensitive chemical. Physical indicators often consist of an alloy designed to melt only after being subjected to 121 °C or 249 °F for 15 minutes. If the alloy melts, the change will be visible.

In addition to these indicators, autoclaves have timers, temperature and pressure gauges that can be viewed from the outside.

There are certain plastics that can withstand repeated temperature cycling greater than the 121 °C or 249 °F required for the autoclaving process. PFA, polypropylene, polysulfone and Noryl are examples.

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 or 249 °F (e.g: F0 = 15 min.). Since exact temperature control is difficult, the temperature is monitored, and the sterilization time adjusted accordingly.

Chemiclave

Unlike the humid environment produced by conventional steam, the unsaturated chemical vapor method is a low-humidity process. No time-consuming drying phase is needed, because nothing gets wet. The heat-up time is shorter than for most steam sterilizers, and the heaters stay on between cycles to minimize warm-up time and increase the instrument turnover

10.Write down principle & application of Flame photo meter?

Flame photometry is an atomic emission method for the routine detection of metal salts, principally Na, K, Li, Ca, and Ba. Quantitative analysis of these species is performed by measuring the flame emission of solutions containing the metal salts. Solutions are aspirated into the flame. The hot flame evaporates the solvent, atomizes the metal, and excites a valence electron to an upper state. Light is emitted at characteristic wavelengths for each metal as the electron returns to the ground state. Optical filters are used to select the emission wavelength monitored for the analyte species. Comparison of emission intensities of unknowns to either that of standard solutions, or to those of an internal standard, allows quantitative analysis of the analyte metal in the sample solution.

Flame photometry is a simple, relatively inexpensive, high sample throughput method used for clinical, biological, and environmental analysis. The low temperature of the natural gas and air flame, compared to other excitation methods such as arcs, sparks, and rare gas plasmas, limit the method to easily ionized metals. Since the temperature isn't high enough to excite transition metals, the method is selective toward detection of alkali and alkali earth metals. On the other hand, the low temperatures renders this method susceptible to certain disadvantages, most of them related to interference and the stability (or lack thereof) of the flame and aspiration conditions. Fuel and oxidant flow rates and purity, aspiration rates, solution viscosity, concomitants in the samples, etc affect these. It is therefore very important to measure the emission of the standard and unknown solutions under conditions that are as nearly identical as possible.

This experiment will serve as an introduction to sodium analysis by flame emission photometry and will demonstrate the effects of cleanliness and solution viscosity on the observed emission intensity readings. The instrument is calibrated with a series of standard solutions that cover the

range of concentrations expected of the samples. Standard calibrations are commonly used in instrumental analysis. They are useful when sample concentrations may vary by several orders of magnitude and when the value of the analyte must be known with a high degree of accuracy. This experiment does not produce hazardous waste.

Procedure

Consult your Teaching Assistant for operating instructions for the Buck PFP-7 Flame Photometer. Allow a sufficient warm-up period. Be sure to aspirate deionized-distilled water between samples to clean out the sample tube and aspirator. Sodium is ubiquitous. It is imperative that you use scrupulously cleaned glassware to obtain good results.

Standard Preparations

Prepare sodium chloride standard solutions by volumetric dilution of the stock solution. The following approximate concentrations should be made: 5, 10, 25, 50, 75, and 100 g/mL as Na. Be sure to use clean methods. Use ultra-pure deionized-distilled water to clean your glassware and for dilution of the 1000 g/mL standard. Prepare these standards in scrupulously clean volumetric glassware and transfer the solutions to plastic bottles. Glass often is made from high sodium glass. Allowing extremely high or low pH solutions to stand in glass could alter the sodium concentrations in solution. Prepare 25 g/mL Na solutions in other solvents, 10% Ethanol, 50% Ethanol, 50% Glycerin. Standard solutions may be pre-prepared by the laboratory instructor or may be made up as a class or group project.

Unknown Preparation

Obtain a sodium unknown from your instructor in a scrupulously clean 50 mL volumetric flask. Dilute to the mark with distilled water.

Instrument Calibration

Set the readout to zero using distilled water as a blank. Set the peak reading according to the instrument instructions using the most concentrated sodium solution (100 g/mL). Measure the emission intensity of each of the remaining sodium standard solutions, and of the sodium unknown solution. Check for accuracy and repeatability by measuring the standards several times. Be sure to aspirate deionized distilled water between measurements.

A photoelectric flame photometer is a device used in inorganic chemical analysis to determine the concentration of certain metal ions, among them sodium, potassium, lithium, and calcium.

In principle, it is a controlled flame test with the intensity of the flame colour quantified by photoelectric circuitry. The sample is introduced to the flame at a constant rate. Filters select which colours the photometer detects and exclude the influence of other ions. Before use, the device requires calibration with a series of standard solutions of the ion to be tested.

Flame photometry is crude but cheap compared to flame emission spectroscopy, where the emitted light is analysed with a monochromator. Its status is similar to that of the colorimeter (which uses filters) compared to the spectrophotometer (which uses a monochromator)

11. Write down principle & application of SDS-PAGE?

INTRODUCTION

Electrophoresis is the migration of charged molecules in solution in response to an electric field. Their rate of migration depends on the strength of the field; on the nett charge, size and shape of the molecules and also on the ionic strength, viscosity and temperature of the medium in which the molecules are moving. As an analytical tool, electrophoresis is simple, rapid and highly sensitive. It is used analytically to study the properties of a single charged species, and as a separation technique.

SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis, is a technique widely used in biochemistry, forensics, genetics and molecular biology to separate proteins according to their electrophoretic mobility (a function of length of polypeptide chain or molecular weight as well as higher order protein folding, posttranslational modifications and other factors).The SDS gel electrophoresis of samples having identical charge to mass ratios results in fractionation by size and is probably the world's most widely used biochemical method

Picture of an SDS-PAGE. The molecular marker is in the left lane

Procedure

The solution of proteins to be analyzed is first mixed with SDS, an anionic detergent which denatures secondary and non–disulfide–linked tertiary structures, and applies a negative charge to each protein in proportion to its mass.[1] [2] [3] Without SDS, different proteins with similar molecular weights would migrate differently due to differences in mass charge ratio, as each protein has an isoelectric point and molecular weight particular to its primary structure. This is known as Native PAGE. Adding SDS solves this problem, as it binds to and unfolds the protein, giving a near uniform negative charge along the length of the polypeptide.

SDS binds in a ratio of approximately 1.4 g SDS per 1.0 g protein (although binding ratios can vary from 1.1-2.2 g SDS/g protein), giving an approximately uniform mass:charge ratio for most proteins, so that the distance of migration through the gel can be assumed to be directly related to only the size of the protein. A tracking dye may be added to the protein solution to allow the experimenter to track the progress of the protein solution through the gel during the electrophoretic run.

Chemical ingredients and its roles

''Polyacrylamide gel (PAGE)'' had been known as a potential embedding medium for sectioning tissues as early as 1954. Two independent groups: Davis and Raymond, employed PAG in electrophoresis in 1959.[4] [5] It possesses several electrophoretically desirable features that made it a versatile medium. PAGE separates protein molecules according to both size and charge. It is a synthetic gel, thermo-stable, transparent, strong, relatively chemically inert, can be prepared with a wide range of average pore sizes [6]. The pore size of a gel is determined by two factors, the total amount of acrylamide present (%T) (T = Total acrylamide-bisacrylamide monomer concentration) and the amount of cross-linker (%C) (C = Crosslinker concentration). Pore size decreases with increasing %T, with cross-linking, 5%C gives the smallest pore size. Any increase or decrease in %C increases the pore size (parabolic function). This appears to be because of nonhomogeneous bundling of strands in the gel.

This gel material can also withstand high voltage gradients, feasible to various staining and destaining procedures and can be digested to extract separated fractions or dried for autoradiography and permanent recording. DISC electrophoresis utilizes gels of different pore sizes. [7] [8] The name DISC was derived from the discontinuities in the electrophoretic matrix and coincidentally from the discoid shape of the separated zones of ions. There are two layers of gel, namely stacking or spacer gel, and resolving or separating gel.

Transmission-Electron Microscopic image of a polyacrylamide gel. The pore size of a gel is determined by the total amount of monomer present (%T) and the amount of cross-linker (%C).

Stacking gel

The stacking gel is a large pore polyacrylamide gel (4%T). This gel is prepared with Tris/HCl buffer pH 6.8 of about 2 pH units lower than that of electrophoresis buffer (Tris/Glycine). These conditions provide an environment for Kohlrausch reactions determining molar conductivity, as a result, SDS-coated proteins are concentrated to several fold and a thin starting zone of the order of 19 μm is achieved in a few minutes. This gel is cast over the resolving gel. The height of the stacking gel region is always maintained more than double the height and the volume of the sample to be applied.

Resolving gel

The resolving gel is a small pore polyacrylamide gel (3 - 30% acrylamide monomer) typically made using a pH 8.8 Tris/HCl buffer. In the resolving gel, macromolecules separate according to their size. Resolving gels have an optimal range of separation that is based on the percent of monomer present in the polymerization reaction; for example an 8%, 10% and 12% resolving gel can effectively used for separating proteins between, 24 – 205 kDa, 14-205 kDa, and 14-66 kDa proteins, respectively (see: SDS gradient gel electrophoresis of proteins).

Chemical ingredients

Tris (tris (hydroxy methyl) aminomethane) (C4H11NO3; mW: 121.14). It has been used as a buffer because it is an innocuous substance to most proteins. Its pKa is 8.3 at 20 °C, making it a very satisfactory buffer in the pH range from roughly 7 to 9.

Glycine (Amino Acetic Acid) (C2H5NO2; mW: 75.07). Glycine has been used as the source of trailing ion or slow ion because its pKa is 9.69 and mobility of glycinate are such that the effective mobility can be set at a value below that of the slowest known proteins of net negative charge in the pH range. The minimum pH of this range is approximately 8.0.

Acrylamide (C3H5NO; mW: 71.08). It is a white crystalline powder. While dissolving in water, autopolymerisation of acrylamide takes place. It is a slow spontaneous process by which acrylamide molecules join together by head on tail fashion. But in presence of free radicals generating system, acrylamide monomers are activated into a free-radical state. These activated monomers polymerise quickly and form long chain polymers. This kind of reaction is known as Vinyl addition polymerisation. A solution of these polymer chains becomes viscous but does not form a gel, because the chains simply slide over one another. Gel formation requires hooking various chains together. Acrylamide is a neurotoxin. It is also essential to store acrylamide in a cool dark and dry place to reduce autopolymerisation and hydrolysis.

Bisacrylamide (N,N'-Methylenebisacrylamide) (C7H10N2O2; mW: 154.17). Bisacrylamide is the most frequently used cross linking agent for poly acrylamide gels. Chemically it is thought of having two-acrylamide molecules coupled head to head at their non-reactive ends.

Sodium Dodecyl Sulfate (SDS) (C12H25NaO4S; mW: 288.38). SDS is the most common dissociating agent used to denature native proteins to individual polypeptides. When a protein mixture is heated to 100 °C in presence of SDS, the detergent wraps around the polypeptide backbone. It binds to polypeptides in a constant weight ratio of 1.4 g/g of polypeptide. In this process, the intrinsic charges of polypeptides becomes negligible when compared to the negative charges contributed by SDS. Thus polypeptides after treatment becomes a rod like structure possessing a uniform charge density, that is same net negative charge per unit length. Mobilities of these proteins will be a linear function of the logarithms of their molecular weights.

Ammonium persulfate (APS) (N2H8S2O8; mW: 228.2). APS is an initiator for gel formation.

TEMED (N, N, N', N'-tetramethylethylenediamine) (C6H16N2; mW: 116.21). Chemical polymerisation of acrylamide gel is used for SDS-PAGE. It can be initiated by ammonium persulfate and the quaternary amine, N,N,N',N'-tetramethylethylenediamine (TEMED). The rate of polymerisation and the properties of the resulting gel depends on the concentration of APS and TEMED. Increasing the amount of APS and TEMED results in a decrease in the average polymer chain length, an increase in gel turbidity and a decrease in gel elasticity. Decreasing the amount of initiators shows the reverse effect. The lowest catalysts concentrations that will allow polymerisation in the optimal period of time should be used. APS and TEMED are used, approximately in equimolar concentrations in the range of 1 to 10 mM.

Chemicals for processing and visualization

The following chemicals are used for processing of the gel and the protein samples visualized in it:

Bromophenol blue (BPB) (3',3",5',5" tetrabromophenolsulfonphthalein) (C19H10Br4O5S; mW: 669.99). BPB is the universal marker dye. Proteins and nucleic acids are mostly colourless. When they are subjected to electrophoresis, it is important to stop the run before they run off the gel. BPB is the most commonly employed tracking dye, because it is viable in alkali and neutral pH, it is a small molecule, it is ionisable and it is negatively charged above pH 4.6 and hence moves towards the anode. Being a small molecule it moves ahead of most proteins and nucleic acids. As it reaches the anodic end of the electrophoresis medium electrophoresis is stopped. It can bind with proteins weakly and give blue colour.

Glycerol (C3H8O3; mW: 92.09). It is a preservative and a weighing agent. Addition of glycerol (20-30 or 50%) is often recommended for the storage of enzymes. Glycerol maintains the protein solution at very low temperature, without freezing. It also helps to weigh down the sample into the wells without being spread while loading.

Coomassie Brilliant Blue (CBB)(C45H44N3NaO7S2; mW: 825.97). CBB is the most popular protein stain. It is an anionic dye, which binds with proteins non-specifically. The structure of CBB is predominantly non-polar. So is usually used (0.025%) in methanolic solution (40%) and acetic acid (7%). Proteins in the gel are fixed by acetic acid and simultaneously stained. The excess dye incorporated in the gel can be removed by destaining with the same solution but without the dye. The proteins are detected as blue bands on a clear background. As SDS is also anionic, it may interfere with staining process. Therefore, large volume of staining solution is recommended, approximately ten times the volume of the gel.

Butanol (C4H10O; mW: 74.12). Water saturated butanol is used as an overlay solution on the resolving gel.

2-Mercaptoethanol (HS-CH2CH2OH; mW: 78.13). 2-Mercaptoethanol is a reducing agent used to disrupt disulfide bonds to ensure the protein is fully denatured before loading on the gel; ensuring the protein runs uniformly.

Reducing SDS-PAGE

Besides the addition of SDS, proteins may optionally be briefly heated to near boiling in the presence of a reducing agent, such as dithiothreitol (DTT) or 2-mercaptoethanol (beta-mercaptoethanol/BME), which further denatures the proteins by reducing disulfide linkages, thus overcoming some forms of tertiary protein folding, and breaking up quaternary protein structure (oligomeric subunits). This is known as reducing SDS-PAGE, and is most commonly used. Non-reducing SDS-PAGE (no boiling and no reducing agent) may be used when native structure is important in further analysis (e.g. enzyme activity, shown by the use of zymograms). For example, quantitative preparative native continuous polyacrylamide gel electrophoresis (QPNC-PAGE) is a new method for separating native metalloproteins in complex biological matrices.

[edit] Electrophoresis and staining

Two SDS-PAGE-gels after a completed run

The denatured proteins are subsequently applied to one end of a layer of polyacrylamide gel submerged in a suitable buffer. An electric current is applied across the gel, causing the negatively-charged proteins to migrate across the gel towards the anode. Depending on their size, each protein will move differently through the gel matrix: short proteins will more easily fit through the pores in the gel, while larger ones will have more difficulty (they encounter more resistance). After a set amount of time (usually a few hours- though this depends on the voltage applied across the gel; higher voltages run faster but tend to produce somewhat poorer resolution), the proteins will have differentially migrated based on their size; smaller proteins will have traveled farther down the gel, while larger ones will have remained closer to the point of origin. Therefore, proteins may be separated roughly according to size (and therefore, molecular weight). Following electrophoresis, the gel may be stained (most commonly with Coomassie Brilliant Blue or silver stain), allowing visualisation of the separated proteins, or processed further (e.g. Western blot). After staining, different proteins will appear as distinct bands within the gel. It is common to run molecular markers of known molecular weight in a separate lane in the gel, in order to calibrate the gel and determine the weight of unknown proteins by comparing the distance traveled relative to the marker. The gel is actually formed because the acrylamide solution contains a small amount, generally about 1 part in 35 of bisacrylamide, which can form cross-links between two polyacrylamide molecules. The ratio of acrylamide to bisacrylamide can be varied for special purposes. The acrylamide concentration of the gel can also be varied, generally in the range from 5% to 25%. Lower percentage gels are better for resolving very high molecular weight proteins, while much higher percentages are needed to resolve smaller proteins. Determining how much of the various solutions to mix together to make gels of particular acrylamide concentration can be done on line

Gel electrophoresis is usually the first choice as an assay of protein purity due to its reliability and ease. The presence of SDS and the denaturing step causes proteins to be separated solely based on size. False negatives and positives are possible. A comigrating contaminant can appear as the same band as the desired protein. This comigration could also cause a protein to run at a different position or to not be able to penetrate the gel. This is why it is important to stain the entire gel including the stacking section. Coomassie Brilliant Blue will also bind with less affinity to glycoproteins and fibrous proteins, which interferes with quantification.

Silver staining

Silver stained SDS Polyacrylamide gels

.

In the 14th century the silver staining technique was developed for colouring the surface of glass. It has been used extensively for this purpose since the 16th century. The colour produced by the early silver stains ranged between light yellow and an orange-red. Camillo Golgi perfected the silver staining for the study of the nervous system. Golgi's method stains a limited number of cells at random in their entirety. The exact chemical mechanism by which this happens is still largely unknown.[9] Silver staining was introduced by Kerenyi and Gallyas as a sensitive procedure to detect trace amounts of proteins in gels.[10] The technique has been extended to the study of other biological macromolecules that have been separated in a variety of supports.[11]

Classical Coomassie Brilliant Blue staining can usually detect a 50ng protein band, Silver staining increases the sensitivity typically 50 times. Many variables can influence the colour intensity and every protein has its own staining characteristics; clean glasware, pure reagents and water of highest purity are the key points to successful staining.[12]

Buffer systems

Postulated migration of proteins in a Laemmli gel system A: Stacking gel, B: Resolving gel, o: sample application c: discontinuities in the buffer and electrophoretic matrix

Most protein separations are performed using a "discontinuous" buffer system that significantly enhances the sharpness of the bands within the gel. During electrophoresis in a discontinuous gel system, an ion gradient is formed in the early stage of electrophoresis that causes all of the proteins to focus into a single sharp band. This occurs in a region of the gel that has larger pores so that the gel matrix does not retard the migration during the focusing or "stacking" event. Negative ions from the buffer in the tank then "outrun" the SDS-covered protein "stack" and eliminate the ion gradient so that the proteins subsequently separate by the sieving action in the lower, "resolving" region of the gel.

Many people continue to use a tris-glycine or "Laemmli" buffering system that stacks at a pH of 6.8 and resolves at a pH of ~8.3-9.0. These pHs promote disulfide bond formation between cysteine residues in the proteins, especially when they are present at high concentrations because the pKa of cysteine ranges from 8-9 and because reducing agent present in the loading buffer doesn't co-migrate with the proteins. Recent advances in buffering technology alleviate this problem by resolving the proteins at a pH well below the pKa of cysteine (e.g., bis-tris, pH 6.5) and include reducing agents (e.g. sodium bisulfite) that move into the gel ahead of the proteins to maintain a reducing environment. An additional benefit of using buffers with lower pHs is that the acrylamide gel is more stable so the gels can be stored for long periods of time before use. [13]

[14]

SDS gradient gel electrophoresis of proteins

Migration of proteins in SDS gels of varying acrylamide concentrations (%T). The migration of nine proteins ranging from 94 kDa to 14.4 kDa is shown. Stacking and unstacking occurs continously in the gel, for every protein at a different gel concentration. The dotted line indicates the discountinuity at the Gly¯/Cl¯ moving boundary. Proteins between the fast leading electrolyte and the slow trailing electrolyte are not diluted by diffusion.

As voltage is applied, the anions (and negatively charged sample molecules) migrate toward the positive electrode in the lower chamber, the leading ion is Cl¯ ( high mobility and high concentration); glycinate is the trailing ion (low mobility and low concentration). SDS-protein particles do not migrate freely at the border between the Cl¯ of the gel buffer and the Gly¯ of the cathode buffer. Friedrich Kohlrausch found that Ohm's law also applies to dissolved electrolytes. Because of the voltage drop between the Cl- and Glycine-buffers, proteins are compressed (stacked) into micrometer thin layers. [15] The boundary moves through a pore gradient and the protein stack gradually disperses due to an frictional resistance increase of the gel matrix. Stacking and unstacking occurs continuously in the gradient gel, for every protein at a different position. For a complete protein unstacking the polyacrylamide-gel concentration must exceed 16% T. The two-gel system of "Laemmli" is a simple gradient gel. The pH discontinuity of the

buffers is of no significance for the separation quality, and a "stacking-gel" with a different pH is not needed.

Support Matrices

Generally the sample is run in a support matrix such as paper, cellulose acetate, starch gel, agarose or polyacrylamide gel. The matrix inhibits convective mixing caused by heating and provides a record of the electrophoretic run: at the end of the run, the matrix can be stained and used for scanning, autoradiography or storage.

In addition, the most commonly used support matrices - agarose and polyacrylamide - provide a means of separating molecules by size, in that they are porous gels. A porous gel may act as a sieve by retarding, or in some cases completely obstructing, the movement of large macromolecules while allowing smaller molecules to migrate freely. Because dilute agarose gels are generally more rigid and easy to handle than polyacrylamide of the same concentration, agarose is used to separate larger macromolecules such as nucleic acids, large proteins and protein complexes. Polyacrylamide, which is easy to handle and to make at higher concentrations, is used to separate most proteins and small oligonucleotides that require a small gel pore size for retardation.

Separation of Proteins and Nucleic Acids

Proteins are amphoteric compounds; their nett charge therefore is determined by the pH of the medium in which they are suspended. In a solution with a pH above its isoelectric point, a protein has a nett negative charge and migrates towards the anode in an electrical field. Below its isoelectric point, the protein is positively charged and migrates towards the cathode. The nett charge carried by a protein is in addition independent of its size - ie: the charge carried per unit mass (or length, given proteins and nucleic acids are linear macromolecules) of molecule differs from protein to protein. At a given pH therefore, and under non-denaturing conditions, the electrophoretic separation of proteins is determined by both size and charge of the molecules.

Nucleic acids however, remain negative at any pH used for electrophoresis and in addition carry a fixed negative charge per unit length of molecule, provided by the PO4 group of each nucleotide of the the nucleic acid. Electrophoretic separation of nucleic acids therefore is strictly according to size.

SDS- PAGE OF PROTEINS

Separation of Proteins under Denaturing conditions

Sodium dodecyl sulphate (SDS) is an anionic detergent which denatures proteins by "wrapping around" the polypeptide backbone - and SDS binds to proteins fairly specifically in a mass ratio of 1.4:1. In so doing, SDS confers a negative charge to the polypeptide in proportion to its length - ie: the denatured polypeptides become "rods" of negative charge cloud with equal

charge or charge densities per unit length. It is usually necessary to reduce disulphide bridges in proteins before they adopt the random-coil configuration necessary for separation by size: this is done with 2- mercaptoethanol or dithiothreitol. In denaturing SDS-PAGE separations therefore, migration is determined not by intrinsic electrical charge of the polypeptide, but by molecular weight.

Determination of Molecular Weight

This is done by SDS-PAGE of proteins - or PAGE or agarose gel electrophoresis of nucleic acids - of known molecular weight along with the protein or nucleic acid to be characterised. A linear relationship exists between the logarithm of the molecular weight of an SDS-denatured polypeptide, or native nucleic acid, and its Rf. The Rf is calculated as the ratio of the distance migrated by the molecule to that migrated by a marker dye-front. A simple way of determining relative molecular weight by electrophoresis (Mr) is to plot a standard curve of distance migrated vs. log10MW for known samples, and read off the logMr of the sample after measuring distance migrated on the same gel.

12.. Write down principle & application of Agarose gel electrophoresis?

Agarose gel electrophoresis

Agarose gel electrophoresis is a method used in biochemistry and molecular biology to separate DNA, or RNA molecules by size. This is achieved by moving negatively charged nucleic acid molecules through an agarose matrix with an electric field (electrophoresis). Shorter molecules move faster and migrate farther than longer ones.[1]

Applications

Estimation of the size of DNA molecules following restriction enzyme digestion, e.g. in restriction mapping of cloned DNA.

Analysis of PCR products, e.g. in molecular genetic diagnosis or genetic fingerprinting Separation of restricted genomic DNA prior to Southern transfer, or of RNA prior to

Northern transfer.

The advantages are that the gel is easily poured, does not denature the samples. The samples can also be recovered.

The disadvantages are that gels can melt during electrophoresis, the buffer can become exhausted, and different forms of genetic material may run in unpredictable forms.

after you finish the experiment and you decide to keep the results store the gel in a plastic bag and in a refrigerater.

Factors affecting migration

The most important factor is the length of the DNA molecule, smaller molecules travel farther. But conformation of the DNA molecule is also a factor. To avoid this problem linear molecules are usually separated, usually DNA fragments from a restriction digest, linear DNA PCR products, or RNAs.

Increasing the agarose concentration of a gel reduces the migration speed and enables separation of smaller DNA molecules. The higher the voltage, the faster the DNA moves. But voltage is limited by the fact that it heats and ultimately causes the gel to melt. High voltages also decrease the resolution (above about 5 to 8 V/cm).

Conformations of a DNA plasmid that has not been cut with a restriction enzyme will move with different speeds (slowest to fastest): nicked or open circular, linearised, or supercoiled plasmid.

Visualisation: Ethidium Bromide (EtBr) and dyes

The most common dye used to make DNA or RNA bands visible for agarose gel electrophoresis is ethidium bromide, usually abbreviated as EtBr. It fluoresces under UV light when intercalated into DNA (or RNA). By running DNA through an EtBr-treated gel and visualizing it with UV light, any band containing more than ~20ng DNA becomes distinctly visible. EtBr is a known carcinogen, however, and safer alternatives are available.

SYBR Green I is another dsDNA stain, produced by Invitrogen. It is more expensive, but 25 times more sensitive, and possibly safer than EtBr, though there is no data addressing its mutagenicity or toxicity in humans.[2]

SYBR Safe is a variant of SYBR Green that has been shown to have low enough levels of mutagenicity and toxicity to be deemed nonhazardous waste under U.S. Federal regulations. [3] It has similar sensitivity levels to EtBr,[3] but, like SYBR Green, is significantly more expensive.

Since EtBr stained DNA is not visible in natural light, scientists mix DNA with negatively charged loading buffers before adding the mixture to the gel. Loading buffers are useful because they are visible in natural light (as opposed to UV light for EtBr stained DNA), and they co-sediment with DNA (meaning they move at the same speed as DNA of a certain length). Xylene cyanol and Bromophenol blue are common loading buffers; they run about the same speed as DNA fragments that are 5000 bp and 300 bp in length respectively, but the precise position varies with percentage of the gel. Other less frequently used progress markers are Cresol Red and Orange G which run at about 125 bp and 50 bp.

Percent agarose and resolution limits

Agarose gel electrophoresis can be used for the separation of DNA fragments ranging from 50 base pair to several megabases (millions of bases) using specialized apparatus. The distance between DNA bands of a given length is determined by the percent agarose in the gel. In general lower concentrations of agarose are better for larger molecules because they result in greater separation between bands that are close in size. The disadvantage of higher concentrations is the long run times (sometimes days). Instead high percentage agarose gels should be run with a pulsed field electrophoresis (PFE), or field inversion electrophoresis.

Most agarose gels are made with between 0.7% (good separation or resolution of large 5–10kb DNA fragments) and 2% (good resolution for small 0.2–1kb fragments) agarose dissolved in electrophoresis buffer. Some people go as high as 3% for separating very tiny fragments but a vertical polyacrylamide gel is more appropriate in this case. Low percentage gels are very weak and may break when you try to lift them. High percentage gels are often brittle and do not set evenly. 1% gels are common for many applications.

Buffers

There are a number of buffers used for agarose electrophoresis. The most common being: tris acetate EDTA (TAE), Tris/Borate/EDTA (TBE) and Sodium borate (SB). TAE has the lowest buffering capacity but provides the best resolution for larger DNA. This means a lower voltage and more time, but a better product. SB is relatively new and is ineffective in resolving fragments larger than 5 kbp; However, with its low conductivity, a much higher voltage could be used (up to 35 V/cm), which means a shorter analysis time for routine electrophoresis. As low as one base pair size difference could be resolved in 3% agarose gel with an extremely low conductivity medium (1 mM Lithium borate).[4]

Analysis

After electrophoresis the gel is illuminated with an ultraviolet lamp (usually by placing it on a light box, while using protective gear to limit exposure to ultraviolet radiation) to view the DNA bands. The ethidium bromide fluoresces reddish-orange in the presence of DNA. The DNA band can also be cut out of the gel, and can then be dissolved to retrieve the purified DNA. The gel can then be photographed usually with a digital or polaroid camera. Although the stained nucleic acid fluoresces reddish-orange, images are usually shown in black and white (see figures).

Gel electrophoresis research often takes advantage of software-based image analysis tools, such as

Typical method

Materials

Typically 10-30 μl/sample of the DNA fragments to separate are obtained, as well as a mixture of DNA fragments (usually 10-20) of known size (after processing with DNA size markers either from a commercial source or prepared manually).

Buffer solution , usually TBE buffer or TAE 1.0x, pH 8.0 Agarose

An ultraviolet-fluorescent dye, ethidium bromide, (5.25 mg/ml in H2O). The stock solution be careful handling this.

Alternative dyes may be used, such as SYBR Green.

Nitrile rubber gloves

Latex gloves do not protect well from ethidium bromide

A color marker dye containing a low molecular weight dye such as "bromophenol blue" (to enable tracking the progress of the electrophoresis) and glycerol (to make the DNA solution denser so it will sink into the wells of the gel).

A gel rack A "comb" Power Supply UV lamp or UV lightbox or other method to visualize DNA in the gel

1 2 3

A 1% agarose 'slab' gel prior to UV illumination, behind a perspex UV shield. Only the marker dyes can be seen

The gel with UV illumination, the ethidium bromide stained DNA glows orange

Digital photo of the gel. Lane 1. Commercial DNA Markers (1kbplus), Lane 2. empty, Lane 3. a PCR product of just over 500 bases, Lane 4. Restriction digest showing the a similar fragment cut from a 4.5 kb plasmid vector

Preparation

There are several methods for preparing gels. A common example is shown here. Other methods might differ in the buffering system used, the sample size to be loaded, the total volume of the gel (typically thickness is kept to a constant amount while length and breadth are varied as needed). Most agarose gels used in modern biochemistry and molecular biology are prepared and run horizontally.

1. Make a 1% agarose solution in 100ml TAE, for typical DNA fragments (see figures). A solution of up to 2-4% can be used if you analyze small DNA molecules, and for large molecules, a solution as low as 0.7% can be used.

2. Carefully bring the solution just to the boil to dissolve the agarose, preferably in a microwave oven.

3. Let the solution cool down to about 60 °C at room temperature, or water bath. Stir or swirl the solution while cooling.

Wear gloves from here on, ethidium bromide is a mutagen, for more information on safety see ethidium bromide

1. Add 5 µl ethidium bromide stock (10 mg/ml) per 100 ml gel solution for a final concentration of 0.5 ug/ml. Be very careful when handling the concentrated stock. Some researchers prefer not to add ethidium bromide to the gel itself, instead soaking the gel in an ethidium bromide solution after running.

2. Stir the solution to disperse the ethidium bromide, then pour it into the gel rack. 3. Insert the comb at one side of the gel, about 5-10 mm from the end of the gel. 4. When the gel has cooled down and become solid, carefully remove the comb. The holes

that remain in the gel are the wells or slots. 5. Put the gel, together with the rack, into a tank with TAE. Ethidium bromide at the same

concentration can be added to the buffer. The gel must be completely covered with TAE, with the slots at the end electrode that will have the negative current.

Procedure

After the gel has been prepared, use a micropipette to inject about 2.5 µl of stained DNA (a DNA ladder is also highly recommended). Close the lid of the electrophoresis chamber and apply current (typically 100 V for 30 minutes with 15 ml of gel). The colored dye in the DNA ladder and DNA samples acts as a "front wave" that runs faster than the DNA itself. When the "front wave" approaches the end of the gel, the current is stopped. The DNA is stained with ethidium bromide, and is then visible under ultraviolet light.

1. The agarose gel with three slots/wells (S). 2. Injection of DNA ladder (molecular weight markers) into the first slot. 3. DNA ladder injected. Injection of samples into the second and third slot. 4. A current is applied. The DNA moves toward the positive anode due to the negative

charges on its phosphate backbone. 5. Small DNA strands move fast, large DNA strands move slowly through the gel. The

DNA is not normally visible during this process, so the marker dye is added to the DNA to avoid the DNA being run entirely off the gel. The marker dye has a low molecular

weight, and migrates faster than the DNA, so as long as the marker has not run past the end of the gel, the DNA will still be in the gel.

6. Add the color marker dye to the DNA ladder.

Agarose gel with samples loaded in the slots, before the electrophoresis process

A pattern of DNA-bands under UV light