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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× magnification
2.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 dry
3.What is the area seen through the ocular eyepiece called?
(a) The stage
(b) The objective
(c) The display
(d) The field of view
4.
(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 specimen
6.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.Aseptic
Aseptic 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 B
1. Describe the prefixes for the metric system and their equivalent in meters.
Prefix
Value in Meters
Kilo (km) (kilo = 1,000)
1,000 m
Deci (dm) (deci = 1/10)
0.10 m
Centi (cm) (centi = 1/100)
0.01 m
Milli (mm) (milli = 1/1000)
0.001 m
Nano (nm) (nano = 1/1,000,000,000)
0.000000001 m
Pico (pm) (pico = 1/1,000,000,000,000)
0.000000000001 m
Kilo (kg)
1,000 g
Hecto (hg)
100 g
Deka (dag)
10 g
Gram (g)
1g
Deci (dg)
0.1 g
Centi (cg)
0.01 g
Milli (mg)
0.001 g
Micro (μg)
0.000001 g
Nano (ng)
0.000000001 g
Pico (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 compound
microscope. These are a smear and a wet mount.
Smear
A 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 Mount
A 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 it
A 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
poun
ds of
press
ure.
At
this
temp
eratu
re
satur
ated
stea
m
destr
oys
all
veget
ative
cells
and
endo
spore
s 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
CH3COO- + H+
CH3COOH
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+]
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
1
,
1
[CH3COO-] [H+]
k1
=
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,
[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,
[A-]
[AH]
pH = pka - log
or pH = pka + log
[A-]
[Salt]
or
[AH]
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 C
1. Explain in detailed about the features of various types of microscopes
Bright-Field Microscope
The 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 MICROSCOPE
A 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 heatsensitive 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.
Mem
bran
e
Filte
r
Steri
lizati
on.
A
mem
brane
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 "concentrationcorrected" 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 organicsoluble 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 law
Main 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 spectrophotometer
See 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 components
Diagram 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:
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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:
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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.
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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:
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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 nondestructive, 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:
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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 purgeand-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, Vinj, 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:
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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
Be sure to use clean methods. Use ultra-pure deionized-distilled water to clean your glassware
are 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
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
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 (betamercaptoethanol/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. Nonreducing 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 (QPNCPAGE) 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 cosediment 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 The gel with UV illumination, Digital photo of the gel. Lane
perspex UV shield. Only the the ethidium bromide stained 1. Commercial DNA Markers
DNA glows orange
(1kbplus), Lane 2. empty, Lane
marker dyes can be seen
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.
2.
3.
4.
The agarose gel with three slots/wells (S).
Injection of DNA ladder (molecular weight markers) into the first slot.
DNA ladder injected. Injection of samples into the second and third slot.
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