M. Sc. Sem - II Botany CBO 406Unit - II : Instrumentation

1. Principles and application of light, phase contrast, fluorescence, scanning and transmission electron microscopy.2. Photometry, colorimetry and spectrophotometry, their application.3. Principles and application of gel-filtration, ion exchange and affinity chromatography. Paper chromatography, thin layer and gas chromatography, HPLC.4. Electrophoresis: PAGE, Agarose gel electrophoresis and electro-focusing, Ultracentrifugation: Principles and types.

MICROSCOPY:

Microscopes, a tool used in various health professions, research institutes, and Universities to magnify small objects that are difficult to see with the naked eye. The intension is to let you have a better understanding of the microscopes and their uses. There two kinds of microscopes that you can identify in any laboratory. They are the simple and compound microscopes magnify an object directly, and on the other hand, compound microscopes magnify an object, and show the objects in their reverse position. You can observe that what is left in the image is right in the object when moving the slide in a certain direction.

LIGHT MICROSCOPY:You have learnt above, the types of microscopes that

require visible light to detect small objects as a well-used research tool in biology. Yet, many of you are unaware of the important features available in these microscopes. For you to use light microscope efficiently, you need to understand the basic microscopy: bright field, dark field, in addition, phase contrast, and oil immersion. When you use any of these fields, you should consider the following; Contrast, Focal Plane, Resolution, and Recognition (CFRR) of the sample when you see it. You should also note that the oil immersion objective (1000x) called “wet” objective, and other lower objectives (400x, 100, and 50x) called “dry” objectives. Bright field microscopy: In bright field microscope, light source is from below the stage. Light travels through the specimen, through the objective lens to your eye through the eyepiece. The microscope controls over the intensity and shape of the light to give an image you see. Bright field microscope gives you best images of stained specimens, naturally pigmented, or living photosynthetic organisms. You should know here that bright field microscope is best used for stained specimens. Phase contrast microscopy: Most of the detail of the transparent living cells is detectable in phase contrast microscopy. However, insufficient contrast between structures with similar transparency may occur. You should know that each transparent structure has a tendency to bend light, providing an opportunity to distinguish them. This translates to mean the reduction in appearance of a structure depends on the refractive index. Highly refractive structure, bend light at much greater angle than do structure with low refractive index. Phase contrast is better than bright field microscopy when the specimen is transparent and high magnifications (400x, 1000x) are required. The health professions and in some university programmes use phase contrast microscopy in teaching. Dark field microscopy: Dark filed microscopy is a cheaper alternative to phase contrast microscopy. The resolution and contrast obtained with the dark field is superior to what you will get from phase contrast, it is important for you to know that in dark field, reflected light from particles on the slide passes through combined lenses (i. e. objectives and eyepieces) to your eye. While phase contrast transmits refracted light through specimen on your slide to your eye. To get a dark field effect you need to place an opaque disc underneath the condenser lens of a bright field microscope; so only external light source (from side of stage or above it) that is scattered by the object on the slide reaches the eye. Any time you want to view specimens in liquid sample, dark field (100X) is best. Oil immersion microscopy: In microbiology lab, you use oil immersion microscopy to observe stained smears of mixed bacteria. You use immersion oil designed especially for oil immersion microscopy. Oil immersion lens is essential for viewing individual bacteria or detail of fixed specimens. To use an immersion lens, you first focus the area of specimen to be observed with the high and dry (400x) lens. Next, you place a drop of immersion oil on the cover slip. Click in the immersion lens, and bring the lens

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down the stage nearly touching the cover slip, while looking from the side. Then you focus by moving the lens up away from the slide until you hit the focal plane to see clear image.

TYPES OF MICROSCOPES AND APPLICATIONS: There are many types of microscopes that you can find in the market today. All you need to do is to determine what it is used for. The different types of microscopes and their basic functions are as listed below. Complex Compound Microscope: Light or optical types of complex compound microscopes combines objective, eyepiece lenses, and light condemner lens and magnify the image of small objects. Simple Compound Microscope: A simple compound microscope is similar to complex compound microscope in function. However, the difference is that a simple compound microscope, is most used to magnifying objects generally. It has no light condenser attached beneath the stage as compared to complex compound

microscope. Simple Magnifying Lens: The simplest light microscope is the magnifying lens. It is usually hand held. It magnifies object and usually useful for field work. Stereo / Dissecting Microscope: A stereo or dissecting microscope, combines two objectives lenses, and two eyepieces to view an object. When you use this microscope, you will see three-dimensional images of the object on the stage. Fluorescence Microscopes: Fluorescence microscope is a special microscope that uses fluorescence and phosphorescence lights to view samples and determine their properties. Digital Microscope: A digital microscope has a digital camera attached to it and connected to a computer screen to view the object directly. It has the

advantages of taking the picture of the object as well. Digital Imager Microscope: This imager microscope is a type of digital video capturing microscope you mount on compound microscope, and connect with USB or AV cable to record the activities of mobile specimens.

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PHASE-CONTRAST MICROSCOPY: is an optical-microscopy technique that converts phase shifts in light passing through a transparent specimen to brightness changes in the image. Phase shifts themselves are invisible, but become visible when shown as brightness variations. When light waves travel through a medium other than vacuum, interaction with the medium causes the wave amplitude and phase to change in a manner dependent on properties of the medium. Changes in amplitude (brightness) arise from the scattering and absorption of light, which is often wavelength-

dependent and may give rise to colors. Photographic equipment and the human eye are only sensitive to amplitude variations. Without special arrangements, phase changes are therefore invisible. Yet, phase changes often carry important information. Phase-contrast microscopy is particularly important in biology. It reveals many cellular structures that are not visible with a simpler bright-field microscope, as exemplified in the figure. These structures were made visible to earlier microscopists by staining, but this required additional preparation and killed the cells. The phase-contrast microscope made it possible for biologists to study living cells and how they proliferate through cell division.[1] After its invention in the early 1930s,[2] phase-contrast microscopy proved to be such an advancement in microscopy, that its inventor Frits Zernike was awarded the Nobel prize (physics) in 1953.

Working principle: Dark field and phase contrast microscopies operating principle. The basic principle to making phase changes visible in phase-contrast microscopy is to separate the illuminating (background) light from the specimen-scattered light (which

makes up the foreground details) and to manipulate these differently. The ring-shaped illuminating light (green) that passes the condenser annulus is focused on the specimen by the condenser. Some of the illuminating light is scattered by the specimen (yellow). The remaining light is unaffected by the specimen and forms the background light (red). When observing an unstained biological specimen, the scattered light is weak and typically phase-shifted by−90° (due to both the typical thickness of specimens and the refractive index difference between biological tissue and the surrounding medium) relative to the background light. This leads to the foreground (blue vector) and background (red vector) having nearly the same intensity, resulting in low image contrast. In a phase-contrast microscope, image contrast is increased in two ways: by generating constructive interference between scattered and background light rays in regions of the field of view that contain the specimen, and by reducing the amount of background light that reaches the image plane. First, the background light is phase-shifted by −90° by passing it through a phase-shift ring, which eliminates the phase difference between the background and the scattered light rays. When the light is then focused on the image plane (where a camera or eyepiece is placed), this phase shift causes background and scattered light rays originating from regions of the field of view that contain the sample (i.e., the foreground) to constructively interfere, resulting in an increase in the brightness of these areas compared to regions that do not contain the sample. Finally, the background is dimmed ~70-90% by a gray filter ring—this method maximizes the amount of scattered light generated by the illumination (i.e., background) light, while minimizing the amount of illumination light that reaches the image plane. Some of the scattered light (which illuminates the entire surface of the filter) will be phase-shifted and dimmed by the rings, but to a much lesser extent than the background light (which only illuminates the phase-shift and gray filter rings). The above describes negative phase contrast. In its positive form, the background light is instead phase-shifted by +90°. The background light will thus be 180° out of phase relative to the scattered light. The scattered light will then be subtracted from the

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background light to form an image with a darker foreground and a lighter background. The success of the phase-contrast microscope has led to a number of subsequent phase-imaging methods.

In 1952 Georges Nomarski patented what is today known as differential interference contrast (DIC) microscopy. It enhances contrast by creating artificial shadows, as if the object is illuminated from the side. But DIC microscopy and phase contrast microscopy both use polarized light, which is unsuitable when the object or its container alter polarization. With the growing use of polarizing plastic containers in cell biology, DIC microscopy is increasingly replaced by Hoffman modulation contrast microscopy, invented by Robert Hoffman in 1975. Traditional phase-contrast methods enhance contrast optically, blending brightness and phase information in single image. Since the introduction of the digital camera in the mid-1990s, several new digital phase-imaging methods have been developed, collectively known as quantitative phase-contrast microscopy. These methods digitally create two separate images, an ordinary bright-field image and a so-called phase-shift image. In each image point, the phase-shift image displays the quantified phase shift induced by the object, which is proportional to the optical thickness of the object.

FLUORESCENCE MICROSCOPY:- An upright fluorescence microscope (Olympus BX61) with the fluorescent filter cube turret above the objective lenses, coupled with a digital camera.

Principle: This is an optical microscope that uses fluorescence and phosphorescence instead of, or in addition to, reflection and absorption to study properties of organic or inorganic substances.[1][2] The "fluorescence microscope" refers to any microscope that uses fluorescence to generate an image, whether it is a more simple set up like an epifluorescence microscope, or a more complicated design such as a confocal microscope, which uses optical sectioning to get better resolution of the fluorescent image. On 8 October 2014, the Nobel Prize in Chemistry was awarded to Eric Betzig, William Moerner and Stefan Hell for "the development of super-resolved fluorescence microscopy," which brings "optical microscopy into the nanodimension". The specimen is illuminated with light of a specific wavelength (or wavelengths) which is absorbed by the fluorophores, causing them to emit light of longer wavelengths (i.e., of a different color than the absorbed light). The illumination light is separated from the much weaker emitted fluorescence through the use of a spectral emission filter. Typical components of a fluorescence microscope are a light source (xenon arc lamp or mercury-vapor lamp are common; more advanced forms are high-power LEDs and lasers), the excitation filter, the dichroic mirror (or dichroic beamsplitter), and the emission filter (see figure below). The filters and the dichroic beamsplitter are chosen to match the spectral excitation and emission characteristics of the fluorophore used to label the specimen.[1] In this manner, the distribution of a single fluorophore (color) is imaged at a time. Multi-color images of several types of fluorophores must be composed by combining several single-color images.

Most fluorescence microscopes in use are epifluorescence microscopes, where excitation of the fluorophore and detection of the fluorescence are done through the same light path (i.e. through the objective). These microscopes are widely used in biology and are the basis for more advanced microscope designs, such as the confocal microscope and the total internal reflection fluorescence microscope (TIRF). A sample of herring spermstained with SYBR green in a cuvette illuminated by blue light in an epifluorescence microscope. The SYBR green in the sample binds to the herring sperm DNAand, once bound, fluoresces giving off green light when illuminated by blue light. In order for a sample to be suitable for fluorescence microscopy it must be fluorescent. There are several methods of creating a fluorescent sample; the main techniques are labelling with fluorescent stains or, in the case of biological

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samples, expression of a fluorescent protein. Alternatively the intrinsic fluorescence of a sample (i.e., autofluorescence) can be used.[1] In the life sciences fluorescence microscopy is a powerful tool which allows the specific and sensitive staining of a specimen in order to detect the distribution of proteins or other molecules of interest. As a result, there is a diverse range of techniques for fluorescent staining of biological samples.

Biological fluorescent stains: Many fluorescent stains have been designed for a range of biological molecules. Some of these are small molecules which are intrinsically fluorescent and bind a biological molecule of interest. Major examples of these are nucleic acid stains like DAPI and Hoechst (excited by UV wavelength light) and DRAQ5 and DRAQ7 (optimally excited by red light) which all bind the minor groove of DNA, thus labeling the nuclei of cells. Others are drugs or toxins which bind specific cellular structures and have been derivatised with a fluorescent reporter. A major example of this class of fluorescent stain is phalloidin which is used to stain actin fibres in mammalian cells. There are many fluorescent molecules called fluorophores or fluorochromes such as fluorescein, Alexa Fluors or DyLight 488, which can be chemically linked to a different molecule which binds the target of interest within the sample.

Limitations: Fluorophores lose their ability to fluoresce as they are illuminated in a process called photobleaching. Photobleaching occurs as the fluorescent molecules accumulate chemical damage from the electrons excited during fluorescence. Photobleaching can severely limit the time over which a sample can be observed by fluorescent microscopy. Several techniques exist to reduce photobleaching such as the use of more robust fluorophores, by minimizing illumination, or by using photoprotective scavenger chemicals.Fluorescence microscopy with fluorescent reporter proteins has enabled analysis of live cells by fluorescence microscopy, however cells are susceptible to phototoxicity, particularly with short wavelength light. Furthermore, fluorescent molecules have a tendency to generate reactive chemical species when under illumination which enhances the phototoxic effect. Unlike transmitted and reflected light microscopy techniques fluorescence microscopy only allows observation of the specific structures which have been labeled for fluorescence. For example, observing a tissue sample prepared with a fluorescent DNA stain by fluorescent microscopy only reveals the organization of the DNA within the cells and reveals nothing else about the cell morphologies.

ELECTRON MICROSCOPY: Principles and capacities: The types of signals produced by an SEM include secondary electrons (SE), reflected or back-scattered electrons (BSE), photons of characteristic X-rays and light (cathodoluminescence) (CL), absorbed current (specimen current) and transmitted electrons. Secondary electron detectors are standard equipment in all SEMs, but it is rare that a single machine would have detectors for all other possible signals.

Types of Electron Microscopes:Transmission electron microscope (TEM) is a microscopy technique in which a beam

of electrons is transmitted through an ultra-thin specimen, interacting with the specimen as it passes through it. An image is formed from the interaction of the electrons transmitted through the specimen; the image is magnified and focused onto an imaging device, such as a fluorescent screen, on a layer of photographic film, or to be detected by a sensor such as a charge-coupled device. TEMs are capable of imaging at a significantly higher resolution than light microscopes, owing to the small de Broglie wavelength of electrons. This enables the instrument's user to examine fine detail—even as small as a single column of atoms, which is thousands of times smaller than the smallest resolvable object in a light microscope. TEM forms a major analysis method in a range of scientific fields, in physical, chemical and biological sciences. TEMs find application in cancer research, virology, materials scienceas well as pollution, nanotechnology and semiconductor research. At smaller magnifications TEM image contrast is due to absorption of electrons in the material, due to the thickness and composition of the material. At higher magnifications complex wave interactions modulate the intensity of the image, requiring expert analysis of observed images. Alternate modes of use allow for the TEM to observe modulations in chemical identity, crystal orientation, electronic structure and sample induced electron phase shift as well as the regular absorption based imaging. The first TEM was built by Max Knoll and Ernst Ruska in 1931, with this group developing the first TEM with resolution greater than that of light in 1933 and the first commercial TEM in 1939. In 1986, Ruska was awarded the Nobel Prize in physics for the development of transmission electron microscopy.

The original form of electron microscope, the transmission electron microscope (TEM) uses a high

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voltage electron beam to illuminate the specimen and create an image. The electron beam is produced by an electron gun, commonly fitted with a tungsten filament cathode as the electron source. The electron beam is accelerated by an anode typically at +100 keV (40 to 400 keV) with respect to the cathode, focused by electrostatic and electromagnetic lenses, and transmitted through the specimen that is in part transparent to electrons and in part scatters them out of the beam. When it emerges from the specimen, the electron beam carries information about the structure of the specimen that is magnified by the objective lens system of the microscope. The spatial variation in this information (the "image") may be viewed by projecting the magnified electron image onto a fluorescent viewing screen coated with a phosphor or scintillator material such as zinc sulfide. Alternatively, the image can be photographically recorded by exposing a photographic film or plate directly to the electron beam, or a high-resolution phosphor may be coupled by means of a lens optical system or a fibre optic light-guide to the sensor of a digital camera. The image detected by the digital camera may be displayed on a monitor or computer. The resolution of TEMs is limited primarily by spherical aberration, but a new generation of aberration correctors have been able to partially overcome spherical aberration to increase resolution. Hardware correction of spherical aberration for the high-resolution transmission electron microscopy (HRTEM) has allowed the production of images with resolution below 0.5 angstrom (50 picometres)[1] and magnifications above 50 million times.[10] The ability to determine the positions of atoms within materials has made the HRTEM an important tool for nano-technologies research and development.

Applications: Transmission electron microscopes are often used in electron diffraction mode. The advantages of electron diffraction over X-ray crystallography are that the specimen need not be a single crystal or even a polycrystalline powder, and also that the Fourier transform reconstruction of the object's magnified structure occurs physically and thus avoids the need for solving the phase problem faced by the X-ray crystallographers after obtaining their X-ray diffraction patterns of a single crystal or polycrystalline powder. The major disadvantage of the transmission electron microscope is the need for extremely thin sections of the specimens, typically about 100 nanometers. Biological specimens are typically required to be chemically fixed, dehydrated and embedded in a polymer resin to stabilize them sufficiently to allow ultrathin sectioning. Sections of biological specimens, organic polymers and similar materials may require special treatment with heavy atom labels in order to achieve the required image contrast.

Scanning electron microscope (SEM) is a type of electron microscope that produces images of a sample by scanning it with a focused beam of electrons. The electrons interact with atoms in the sample, producing various signals that contain information about the sample's surface topography and composition. The electron beam is generally scanned in a raster scan pattern, and the beam's position is combined with the detected signal to produce an image. SEM can achieve resolution better than 1 nanometer. Specimens can be observed in high vacuum, in low vacuum, in wet conditions (in environmental SEM), and at a wide range of cryogenic or elevated temperatures. The most common SEM mode is detection of secondary electrons emitted by atoms excited by the electron beam. The number of secondary electrons that can be detected depends, among other things, on specimen topography. By scanning the sample and collecting the secondary electrons that are emitted using a special detector, an image displaying the topography of the surface is created. The SEM produces images by probing the specimen with a focused electron beam that is scanned across a rectangular area of the specimen (raster scanning). When the electron beam interacts with the specimen, it loses energy by a variety of mechanisms. The lost energy is converted into alternative forms such as heat, emission of low-energy secondary electrons and high-energy backscattered electrons, light emission (cathodoluminescence) or X-rayemission, all of which provide signals carrying information about the properties of the specimen surface, such as its topography and composition. The image displayed by an SEM maps the varying intensity of any of these signals into the image in a position corresponding to the position of the beam on the specimen when the signal was generated. In the SEM image of an ant shown below and to the right, the image was constructed from signals produced by a secondary electron detector, the normal or conventional imaging mode in most SEMs.

Generally, the image resolution of an SEM is at least an order of magnitude poorer than that of a TEM. However, because the SEM image relies on surface processes rather than transmission, it is able to image bulk samples up to many centimeters in size and (depending on instrument design and settings) has a great depth of field, and so can produce images that are good representations of the three-dimensional shape of the sample. Another advantage of SEM is its variety called environmental scanning electron microscope (ESEM) can produce images of sufficient quality and resolution with the samples being wet or

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contained in low vacuum or gas. This greatly facilitates imaging biological samples that are unstable in the high vacuum of conventional electron microscopes. Color: In their most common configurations, electron microscopes produce images with a single brightness value per pixel, with the results usually rendered in grayscale.[12] However, often these images are then colorized through the use of feature-detection software, or simply by hand-editing using a graphics editor. This may be done to clarify structure or for aesthetic effect and generally does not add new information about the specimen. In some configurations information about several specimen properties is gathered per pixel, usually by the use of multiple detectors.[14] In SEM, the attributes of topography and material contrast can be obtained by a pair of backscattered electron detectors and such attributes can be superimposed in a single color image by assigning a different primary color to each attribute. [15] Similarly, a combination of backscattered and secondary electron signals can be assigned to different colors and superimposed on a single color micrograph displaying simultaneously the properties of the specimen.

Applications: Some types of detectors used in SEM have analytical capabilities, and can provide several items of data at each pixel. Examples are the Energy-dispersive X-ray spectroscopy(EDS) detectors used in elemental analysis and Cathodoluminescence microscope (CL) systems that analyse the intensity and spectrum of electron-induced luminescence in (for example) geological specimens. In SEM systems using these detectors it is common to color code the signals and superimpose them in a single color image, so that differences in the distribution of the various components of the specimen can be seen clearly and compared. Optionally, the standard secondary electron image can be merged with the one or more compositional channels, so that the specimen's structure and composition can be compared. Such images can be made while maintaining the full integrity of the original signal, which is not modified in any way. Disadvantages of Electron Microscopy: Electron microscopes are expensive to build and maintain, but the capital and running costs of confocal light microscope systems now overlaps with those of basic electron microscopes. The samples largely have to be viewed in vacuum, as the molecules that make up air would scatter the electrons. An exception is the environmental scanning electron microscope, which allows hydrated samples to be viewed in a low-pressure and/or wet environment. Samples of hydrated materials, including almost all biological specimens have to be prepared in various ways to stabilize them, reduce their thickness (ultrathin sectioning) and increase their electron optical contrast (staining). These processes may result in artifacts, but these can usually be identified by comparing the results obtained by using radically different specimen preparation methods.

Reflection electron microscope (REM): This technique is typically coupled with reflection high energy electron diffraction (RHEED) and reflection high-energy loss spectroscopy (RHELS). Another variation is spin-polarized low-energy electron microscopy (SPLEEM), which is used for looking at the microstructure of magnetic domains.

Scanning transmission electron microscope (STEM): The STEMs use of SEM-like beam rastering simplifies annular dark-field imaging, and other analytical techniques, but also means that image data is acquired in serial rather than in parallel fashion. Often TEM can be equipped with the scanning option and then it can function both as TEM and STEM.

PHOTOMETRY:

What is light? … The light what we see is just a small region of electromagnetic spectrum, which human eyes are capable of sense. This region is called as "visible light". Where this light comes from? Due to fusion reaction in the sun. When two hydrogen atoms fused with each other, they form a helium molecule. In this process some mass is removed in the form of energy. This energy is the electromagnetic radiation. This is partly electric and partly magnetic, so it is called as electromagnetic radiation. Why we can see any object and how could we define colors?... When the visible light is reflected from any object and touches the sensors of our eyes, we can see the objects. In the process of reflection- out of multiple colors of the radiation, some colors are absorbed and some are reflected. The color which is reflected is the color what we can see. Why black colored car is hotter than white colored car during summer?... All the frequencies of white light (all colors) are absorbed by the black

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colored car while all the frequencies of white light (all colors) are reflected by white colored car. So, black colored car is hotter than white colored car during summer.

Photometry is the measurement of electromagnetic radiation weighted by the human eye's response. This response changes with wavelength, and to an extent, from person to person. Internationally-agreed standard observer functions are therefore used in order to provide a consistent measurement base for photometry; the two most widely used are the V(λ) function, which applies for photopic vision (typical day-time light levels) and the V'(λ) for scotopic vision (low lighting levels). At intermediate light levels (mesopic or ‘twilight’ levels, such as found on lit roads at night), the CIE system of mesopic photometry is used to provide a smooth transition between these two functions. In photometry, the word 'luminous' is used to indicate that measurements have been made using a detection system (called a photometer) that has a spectral response similar to that of a human eye. The two principal photometric scales maintained at NPL are of luminous intensity and luminous flux. Setting up appropriate geometries permits calibrations of other quantities, such as luminance from luminous intensity standards. NPL has extensive facilities available for the photometric measurement of both sources and detectors, including photometers, luxmeters, luminance meters and colour temperature meters. Services include the calibration of luminous intensity, illuminance, luminance, luminous flux and correlated colour and temperature. Luminance (photometric) and radiance (radiometric), Luminous flux (photometric) and radiant flux (radiometric), Luminous intensity (photometric) and radiant intensity (radiometric) are the units.

PHOTOMETRIC MEASUREMENT TECHNIQUES: Photometric measurement is based on photodetectors, devices (of several types) that produce and electric signal when exposed to light. Simple applications of this technology include switching luminaires on and off based on ambient light conditions, and light meters, used to measure the total amount of light incident on a point. More complex forms of photometric measurement are used frequently within the lighting industry. Spherical photometers can be used to measure the directional luminous flux produced by lamps, and consist of a large-diameter globe with a lamp mounted at its center. A photocell rotates about the lamp in three axes, measuring the output of the lamp from all sides. Luminaires (known to laypersons simply as light fixtures) are tested using

goniophotometers and rotating mirror photometers, which keep the photocell stationary at a sufficient distance that the luminaire can be considered a point source. Rotating mirror photometers use a motorized system of mirrors to reflect light emanating from the luminaire in all directions to the distant photocell; goniophotometers use a rotating 2-axis table to change the orientation of the luminaire with respect to the photocell. In either case, luminous intensity is tabulated from this data and used in lighting design.

Applications: Photometry is the measurement of light, in terms of its perceived brightness to the human

eye. Many units of measure are used for photometric measurements. The important points to note in this unit are: Photometry is the measurement of light as its perceived brightness to the human eye. The human eye is not equally sensitive to all wavelengths of visible light. Photometric and radiometric quantities are the two parallel systems of calorimetry. Watts are units of radiant flux while lumens are units of luminous flux. The watt is a unit of power

COLORIMETRY: The colorimeter is based on Beer-Lambert's law, according to which the absorption of light transmitted through the medium is directly proportional to the medium concentration. In a colorimeter, a beam of light with a specific wavelength is passed through a solution via a series of lenses, which navigate the colored light to the measuring device. This analyses the color compared to an existing standard. A microprocessor then calculates the absorbance or percent transmittance. If the concentration

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of the solution is greater, more light will be absorbed, which can be identified by measuring the difference between the amount of light at its origin and that after passing the solution. In order to determine the concentration of an unknown sample, several sample solutions of a known concentration are first prepared and tested. The concentrations are then plotted on a graph against absorbance, thereby generating a calibration curve. The results of the unknown sample are compared to that of the known sample on the curve to measure the concentration.

Draw and explain the spectrum of light. As we move from left to right side the frequency (vibrations/sec) of light decreases and wavelength (length of a wave) increases. Some time we get line of colors so they are called as line spectra or discontinued spectra or emission spectra. Eg. Hydrogen spectra.

Max plank = Light is having particle nature, Hugense = Light is having wave nature, Einstine = Light is having both (particle and wave) nature. As an atom the smallest part of light is called as photon. It can be imagined as an energy packet. Speed of light = 3.0 x 108m/sec. RGB (red, green and blue) are basic colors and all other are the derivatives of these primary colors.

Colorimetry…. Monochromatic Light: A single colored light which is having a particular frequency is called as monochromatic light (Monochromators): Instruments used for getting a monochromatic light are called as monochromators. For eg. Prism, grating etc. Explain Beer and Lambert law. As a mono chromatic

light is passed through a solution, some intensity is absorbed in the solution. So, difference between the intensity of initial and transmitted light occurs. Beer's Law: As the concentration increases the absorption increase and so intensity of transmitted light decreases exponentially. Lambert's Law: As the thickness (or sometimes taken as cell length) increases the absorption increase and so intensity of transmitted light decreases exponentially. A = a be Where, A = absorbance a = absorptivity; experimentally derived constant for each substrate b = path length of light through sample c = concentration of substance in solution. Absorbance…A = - log T, T = 10**, log T = log (10 ■*bc), log T = -abc, , -log T = -(-abc) = abc, A = -log T = abcAs given in the above figure, as the transmission increases, absorbance decreases exponentially. Deviation in Beer and Lambert’s Law: If a substance is following the beer's law, we get a straight line which is passing from the origin and the slope is "ab". But some time we do not get a straight line. If we are getting much more value of absorbance than desired, than deviation is said as +ve deviation. But if we are getting much less value of absorbance than desired, than deviation is said as -ve deviation.

Beer's Law: … There are two reason for these deviations: Instrumental Errors: The cause of instrumental errors are as follows: Fluctuation in electricity Source of light is weak or malfunction Arrangement of filter/monochromator is not proper Scattering of light is happening inside the instrument Slit

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is not placed properly Outside knob are not working according to inner instrumentation Sensitivity of detector is low or malfunction Chemical Errors: The cause of chemical errors are as follows: Presence of bacteria Solution is cloudy Pigmentation is there in the solution Acid-Base reaction is happening in the solution Association-Dissociation reaction is happening in the solution. Polarization reaction is happening in the solution. If the color of the solution is changing with time. If the absorption is happening due to solvent rather than solute. This law is only applicable to some extent of concentration of the solution only. Below or above that concentration these laws are not applicable.INSTRUMENTATION OF COLORIMETER:Colorimeter is an instrument which compares the amount of light getting through unknown solution and the amount of light getting through a pure solvent.❖ Instrumentation:The source light is passed through filter (to get desired frequency light) and concentrated using lens. Then this light is targeted on a sample (which is kept in cuvette). The light transmits through the sample and the transmitted light is measured by detector. As the intensity of the initial light is known, we can find the difference between - intensity of initial light and intensity of transmitted light.

Applications of colorimeter: First of all, instrument is set to 100% transmission (0% absorption) for cuvette with the solvent only. Then known concentrations of the desired solution is prepared. (For eg. 1 ppm, 2 ppm, 3 ppm...) Then the absorptions related to the known concentrations are noted using the colorimeter. From the readings of absorption, a calibration curve is drawn. (graph: absorption vs concentration). Then the absorption of the known concentration - of known solution is acquired using the colorimeter instrument and from the calibration curve the concentration of the unknown solution could be acquired. Pre-requisite for a solution to be analyzed by colorimeter: The solution must be colored. The solution must not be having any contamination like Bacteria, Cloudy solution or Pigmentation. The solution must be clear (transparent). There must not be any reaction happening in the solution like acid-base, association-dissociation or polarization reaction. The solution must be having a particular concentration. Because lambert and beer laws can be applied only for particular range of concentration. Colorimeters are widely used to monitor the growth of a bacterial or yeast culture. They provide reliable and highly accurate results when used for the assessment of color in bird plumage. They are used to measure and monitor the color in various foods and beverages, including vegetable products and sugar. Certain colorimeters can measure the colors that are used in copy machines, fax machines and printers. Besides being used for basic research in chemistry laboratories, colorimeters have many practical applications such as testing water quality by screening chemicals such as chlorine, fluoride, cyanide, dissolved oxygen, iron, molybdenum, zinc and hydrazine. They are also used to determine the concentrations of plant nutrients such as ammonia, nitrate and phosphorus in soil or hemoglobin in blood.

SPECTROPHOTOMETRY: In chemistry, spectrophotometry is the quantitative measurement of the reflection or transmission properties of a material as a function of wavelength. It is more specific than the general term electromagnetic spectroscopy in that spectrophotometry deals with visible light, near-ultraviolet, and near-infrared, but does not cover time-resolved spectroscopic techniques. Spectrophotometry uses photometers, known as spectrophotometers that can measure a light beam's intensity as a function of its color (wavelength). Important features of spectrophotometers are spectral bandwidth (the range of colors it can transmit through the test sample), the percentage of sample-transmission, the logarithmic range of sample-absorption, and sometimes a percentage of reflectance measurement. A spectrophotometer is commonly used for the measurement of transmittance or reflectance of solutions, transparent or opaque solids, such as polished glass, or gases. However they can also be designed to measure the diffusivity on any of the listed light ranges that usually cover around 200 nm - 2500 nm using different controls and calibrations. Within these ranges of light, calibrations are needed on the machine using standards that vary in type depending on the wavelength of the photometric determination. An example of an experiment in

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which spectrophotometry is used is the determination of the equilibrium constant of a solution. A certain chemical reaction within a solution may occur in a forward and reverse direction where reactants form products and products break down into reactants. At some point, this chemical reaction will reach a point of balance called an equilibrium point. In order to determine the respective concentrations of reactants and products at this point, the light transmittance of the solution can be tested using spectrophotometry. The amount of light that passes through the solution is indicative of the concentration of certain chemicals that do not allow light to pass through. The use of spectrophotometers spans various scientific fields, such as physics, materials science, chemistry, biochemistry, and molecular biology. They are widely used in many industries including semiconductors, laser and optical manufacturing, printing and forensic examination, as well in laboratories for the study of chemical substances. Ultimately, a spectrophotometer is able to determine, depending on the control or calibration, what substances are present in a target and exactly how much through calculations of observed wavelengths.

❖ Difference between filter and monochromators: By filter a specific band of frequency can be acquired while by monochromators a specific ray of light can

be acquired.❖ Wave length selection method: Use other filter than

the color of the solution. Because the color of the solution is the color which is not absorbed by the solution. For eg. In case of CuSO4 solution, use red colored filter. Find the highest absorption using

different filters of different colors. The wavelength recommended for different solutions is given in the S.O.P. (standard operating procedure) manual of the instrument. So use that filter. Spectrophotometry❖ Instrumentation of spectrophotometer: There are two types spectrophometer available:Types: Single beam spectrophotometer & double beam spectrophotometer

Detector, Sample, Detector, Dispersion, Entrance slit, SourceSource of light: The light source must be fulfilling the following requisitions: The light coming from the source must be having proper intensity. The light source must be having all the frequencies of light, so that the required frequency can be acquired. The light source must be stable. It must not change with time. For different region of light, following lamps (light sources) can be used:Filter and Monochromators: From this part, the radiation with only specific wavelength or specific wavelength could be passed. Other radiations are absorbed. Filters allow only a small section of frequency to pass through and all others are absorbed. Filters are used mostly in colorimeter. They are made up of glass or gelatin. By using different filters we can separate different regions of visible light as follows: * Monochromators: Optical devices used for selecting a specific wavelength from a range of frequency are called monochromators. They are of two types: 1. Slit 2. Dispersive element for dispersion of light, prism or grating or both can be used. Sample vessel (Cuvette): In all the spectrophotometric and analysis the absorption of light is measured. So the cell used in the analysis must not absorb the light or the absorption must be minimum. Hence, for different region of spectrum - different material could be used as follows: 1. For UV range: Quartz cuvette, for visible range: glass cuvette, For IR range: NaCl, KBr, nujol cuvette, Detector: The light which is initiated from the source and pass through cuvette is measured by the detector. Absorption or transmission of the light can be measured by detector. Mainly three type of detectors can be used: Photovoltaic Cell (Barrier layer Cell), Photo tubes (Photo emissive tube), Photo multiplier tubes.* Recorder:The measurement of absorption or transmission is recorded in digital form in recorder. The graph of wavelength vs absorption can also be acquired. Figure for photo multiplier is given below: ❖ Photo multiplier Tube: Due to very high amplification in Photo multiplier tube, it is much sensitive than simple photo cell tube. So It is very much used in spectrophotometer. There is a photo cathode, the surface of which is coated with light reflecting material. There are also symmetrically arranged poles which are called as dynode. Each dynode is also having light reflecting material. Which is having more potential than

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Color of visible region wavelength1. Violet 380-470 nm2. Blue 440-490 nm3. Blue (slightly greenish) 490-500 nm4. Green 500-560 nm5. Yellow (slightly greenish) 560-580 nm6. Yellow 580-600 nm7. Orange 600-650 nm8. Red 650-750 nm

EM Region Light Source Wavelength1.

U.V. / near IR region Hydrogen or Deuterium discharge Lamp

10-200 nm2.

Visible region Tungsten Lamp 200-1000 nm3 IR region Nernst glover 1000-10,00,000 nm

sequential cathode? Dynode is set to +ve voltage. So, when light indented on the surface of cathode, a primary electron is generated. These electrons flows to the neighboring dynode whose potential is 50-90 v more than cathode. Moreover each electron genereates 4-5 secondary electrons. This process happens on almost 9 dynode. Thus, amplification happens in the tube and shower of electron (49 = 2.6 x 106) occurs. When this shower of electron is captured by detector, intensity of the light can be known and thus we can have strong signal. To start the tube, 500 to 900 V of electricity is necessary which is applied using no. of batteries in a sequence.

Applications of Spectrophotometry: Spectrophotometry is an important technique used in many biochemical experiments that involve DNA, RNA, and protein isolation, enzyme kinetics and biochemical analyses. A brief explanation of the procedure of spectrophotometry includes comparing the absorbency of a blank sample that does not contain a colored compound to a sample that contains a colored compound. The spectrophotometer is used to measure colored compounds in the visible region of light (between 350 nm and 800 nm), thus it can be used to find more information about the substance being studied. In biochemical experiments, a chemical and/or physical property is chosen and the procedure that is used is specific to that property in order to derive more information about the sample, such as the quantity, purity, enzyme activity, etc. Spectrophotometry is also a helpful procedure for protein purification and can also be used as a method to create optical assays of a compound. Because a spectrophotometer measures the wavelength of a compound through its color, a dye binding substance can be added so that it can undergo a color change and be measured. Spectrophotometers have been developed and improved over decades and have been widely used among chemists. It is considered to be a highly accurate instrument that is also very sensitive and therefore extremely precise, especially in determining color change. This method is also convenient for use in laboratory experiments because it is an inexpensive and relatively simple process. Estimating dissolved organic carbon concentration. Specific Ultraviolet Absorption for metric of aromaticity. Bial's Test for concentration of pentoses.

CHROMATOGRAPHY:

Chromatography is separation and analytical techniques widely used in chemistry and the biological sciences. Most things that occur in nature are a mixture of substances which can only be separated or analysed using any of the techniques known. In this unit (3) you will learn about some of these techniques.

Chromatograpy, firstly introduced by the Russian botanist Michael Iswett is a method for separating the components of a mixture by differential distribution of the components between a stationary phase and mobile (moving) phase. Initially used for the separation of coloured substances from the plants (Greek, Chromos meaning coloured) is now the most extensive technique of separation and purification of coloured/ colourless organic compounds. Separation of two sample components in chromatography is based on their different distribution between two non-miscible phases. The one, the stationary phase, a liquid or solid, is fixed in the system. The other, the mobile phase, a fluid, is streeming through the

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chromatographic system. In gas chromatography the mobile phase is a gas, in liquid chromatography it is a liquid. The molecules of the analytes (mixture to be separated) are distributed between the mobile and the stationary phase. When present in the stationary phase, they are retained, and are not moving through the system. In contrast, they migrate with the velocity, v, of the mobile phase when being there. Due to the different distribution of the particular analytes the mean residence time in the stationary phase differs, too, resulting in a different net migration velocity. This is the principle of chromatographic separation. Separation of two sample components in chromatography is based on their different distribution between two non-miscible phases.

GEL FILTRATION CHROMATOGRAPHY (Size Exclusion Chromatography): Gel filtration chromatography seprarates proteins, peptides, and oligonucleotides on the basis of size. Molecules move through a bed of porous beads, diffusing into the beads to greater or lesser degrees. Smaller molecules diffuse further into the pores of the beads and therefore move through the bed more slowly, while larger molecules enter less or not at all and thus move through the bed more quickly. Both molecular weight and three-dimensional shape contribute to the degree of retention. Gel Filtration Chromatography may be used for analysis of molecular size, for separations of components in a mixture, or for salt removal or buffer exchange from a preparation of macromolecules. Size exclusion chromatography is used for semi-

preparative purifications and various analytical assays. It is a separation technique which takes the advantage of the difference in size and geometry of the molecules. The molecules are separated based on their size. Grant Henry Lathe and Colin R Ruthven was the pioneer of size exclusion chromatography who started this technique for separation of analytes of different size with starch gels as the matrix, later Jerker Porath and Per Flodin introduced dextran gels. Other gel filtration matrices include agarose and polyacrylamide. Note: Unlike ion exchange chromatography, gel filtration does not depend on any chemical interaction with

protein, rather it is based on a physical property of the protein - that being the effective molecular radius (which relates to mass for most globular proteins).

Principle: Size exclusion chromatography (SEC) is the separation of mixtures based on the molecular size (more correctly, their hydrodynamic volume) of the components. Separation is achieved by the differential exclusion or inclusion of solutes as they pass through stationary phase consisting of heteroporous (pores of different sizes) cross linked polymeric gels or beads. The process is based upon different permeation rates of each solute molecule into the interior of gel particles. Size exclusion chromatography involves gentle interactionwith the sample, enabling high retention of biomolecular activity. For the separation of biomolecules in aqueous systems, SEC is referred to as gel filtration chromatography (GFC), while the separation of organic polymers in non-aqueous systems is called gel permeation chromatography (GPC). The basic principle of size exclusion chromatography is quite simple. A column of gel particles or porous matrix is in equilibrium with a suitable mobile phase for the molecules to be separated. Large molecules are completely excluded from the pores will pass through the space in between the gel particles or matrix and will come first in the effluent. Smaller molecules will get distributed in between the mobile phase of in and outside the

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molecular sieve and will then pass through the column at a slower rate, hence appear later in effluent. There are two extremes in the separation profile of a gel filtration column. There is a critical molecular mass (large mass) which will be completely excluded from the gel filtration beads. All solutes in the sample which are equal to, or larger, than this critical size will behave identically: they will all eluted in the excluded volume of the column. There is a critical molecular mass (small mass) which will be completely included within the pores of the gel filtration beads. All solutes in the sample which are equal to, or smaller, than this critical size will behave identically: they will all eluted in the included volume of the column. Solutes between these two ranges of molecular mass will elute between the excluded and included volumes, Thus, while deciding a size exclusion matrix for proteinpurification, included and excluded range should be considered. For example: Sephadex G 75 matrix has fractionation range 3-80. This tells that the matrix has included volume range 3 kDa and excluded volume range 80kDa. If protein of interest and impurities both are close to 80 kDa or above they are likely to co-elute in excluded volume. Thus purification will not work. Now you can think what is the use of a size exclusion matrix Sephadax G25 (range 1- 5kDa)? This is generally used for desalting as all proteins are above 5kDa and comes in excluded volume and salts are eluted late in included volume. In gel filtration the resolution is a function of column length (the longer the better). However, one drawback is related to the maximum sample volume which can be loaded. The larger the volume of sample loaded, the more the overlap between separated peaks. Generally speaking, the sample size one can load is limited to about 3-5% of the total column volume. Thus, gel filtration is best saved for the end stages of a purification, when the sample can be readily concentrated to a small volume. Gel filtration can also be used to remove salts from the sample, due to its ability to separate "small" from "large" components. Finally, gel filtration can be among the most "gentle" purification methods due to the lack of chemical interaction with the resin.

Applications: Purification. Desalting. Protein-ligand binding studies. Protein folding studies. Concentration of sample. Copolymerisation studies. Relative molecular mass determination.

ION EXCHANGE CHROMATOGRAPHY (ION CHROMATOGRAPHY): is a chromatography process that separates ions and polar molecules based on their affinity to the ion exchanger. It works on almost any kind of charged molecule—including large proteins, small nucleotides, and amino acids. It is often used in protein purification, water analysis, and quality control.[citation needed] The water-soluble and charged molecules such as proteins, amino acids, and peptides bind to moieties which are oppositely charged by forming ionic bonds to the insoluble stationary phase. [1] The equilibrated stationary phase consists of an ionizable functional group where the targeted molecules of a mixture to be separated and quantified can bind while passing through the column—a cationic stationary phase is used to separate anions and an anionic stationary phase is used to separate cations. Cation exchange chromatography is used when the desired molecules to separate are cations and anion exchange chromatography is used to separate anions.[2] The bound molecules then can be eluted and collected using an eluant which contains anions and cations by running higher concentration of ions through the column or changing pH of the column. One of the primary advantages for the use of ion chromatography is only one interaction involved during the separation as opposed to other separation techniques; therefore, ion chromatography may have higher matrix tolerance. However, there are also disadvantages involved when performing ion-exchange chromatography, such as constant evolution with the technique which leads to the inconsistency from column to column.

Principle: Ion-exchange chromatography separates molecules based on their respective charged

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groups. Ion-exchange chromatography retains analyte molecules on the column based on coulombic (ionic) interactions. Essentially, molecules undergo electrostatic interactions with opposite charges on the stationary phase matrix. The stationary phase consists of an immobile matrix that contains charged ionizable functional groups or ligands. [10] The stationary phase surface displays ionic functional groups (R-X) that interact with analyte ions of opposite charge. To achieve electroneutrality, these inert charges couple with exchangeable counterions in the solution. Ionizable molecules that are to be purified compete with these exchangeable counterions for binding to the immobilized charges on the stationary phase. These ionizable molecules are retained or eluted based on their charge. Initially, molecules that do not bind or bind weakly to the stationary phase are first to wash away. Altered conditions are needed for the elution of the molecules that bind to the stationary phase. The concentration of the exchangeable counterions, which competes with the molecules for binding, can be increased or the pH can be changed. A change in pH affects the charge on the particular molecules and, therefore, alters binding. The molecules then start eluting out based on the changes in their charges from the adjustments. Further such adjustments can be used to release the protein of interest. Additionally, concentration of counterions can be gradually varied to separate ionized molecules. This type of elution is called gradient elution. On the other hand, step elution can be used in which the concentration of counterions are varied in one step. [11] This type of chromatography is further subdivided into cation exchange chromatography and anion-exchange chromatography. Positively charged molecules bind to anion exchange resins while negatively charged molecules bind to cation exchange resins. 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, thus retention time. The ion chromatogram shows a typical chromatogram obtained with an anion exchange column.

Typical technique: 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. Equilibration of the stationary phase is needed in order to obtain the desired charge of the column. If the column is not properly equilibrated the desired molecule may not bind strongly to the column. 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 displaces the analyte ions from the stationary phase. For example, in cation exchange chromatography, the positively charged analyte can be displaced by adding positively charged sodium ions. The analytes of interest must then be detected by some means, typically by conductivity or UV/visible light absorbance. Control an IC system usually requires a chromatography data system (CDS). In addition to IC systems, some of these CDSs can also control gas chromatography (GC) and HPLC.

Applications: Clinical utility: A use of ion chromatography can be seen in the argentation ion chromatography.[citation needed] Usually silver and compounds containing acetylenic and ethylenic bonds have very weak interactions. This phenomenon has been widely tested on olefin compounds. The ion complexes the olefins make with silver ions are weak and made based on the overlapping of pi, sigma, and d orbitals and available electrons therefore cause no real changes in the double bond. This behavior was manipulated to separate lipids, mainly fatty acids from mixtures in to fractions with differing number of double bonds using silver ions. Industrial applications: Since 1975 ion chromatography has been widely used in many branches of industry. The main beneficial advantages are reliability, very good accuracy and precision, high selectivity, high speed, high separation efficiency, and low cost of consumables. The most significant development related to ion chromatography are new sample preparation methods; improving the speed and selectivity of analytes separation; lowering of limits of detection and limits of quantification; extending the scope of applications; development of new standard methods; miniaturization and extending the scope of the analysis of a new group of substances. Drug development: An ion chromatography system used to detect and measure cations such as sodium, ammonium and potassium in Expectorant Cough Formulations. There has been a growing interest in the application of IC in the analysis of pharmaceutical drugs. Detection of sugar and sugar alcohol in such formulations through IC has been done due to these polar groups getting resolved in ion column.

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AFFINITY CHROMATOGRAPHY: is a method of separating biochemical mixtures based on a highly specific interaction such as that between antigen and antibody, enzyme and substrate, or receptor and ligand.

Principle: The stationary phase is typically a gel matrix, often of agarose; a linear sugar molecule derived from algae. Usually the starting point is an undefined heterogeneous group of molecules in solution, such as a cell lysate, growth medium or blood serum. The molecule of interest will have a well-known and defined property, and can be exploited during the affinity purification process. The process itself can be thought of as an entrapment, with the target molecule becoming trapped on a solid or stationary phase or medium. The other molecules in the mobile phase will not become trapped as they do not possess this property. The stationary phase can then be removed from the mixture, washed and the target molecule released from the entrapment in a process known as elution. Possibly the most common use of affinity chromatography is for the purification of recombinant proteins.

Batch chromatography: Binding to the solid phase may be achieved by column chromatography whereby the solid medium is packed onto a column, the initial mixture run through the column to allow setting, a wash buffer run through the column and the elution buffer subsequently applied to the column and collected. These steps are usually done at ambient pressure. Alternatively, binding may be achieved using a batch treatment, for example, by adding the initial mixture to the solid phase in a vessel, mixing, separating the solid phase, removing the liquid phase, washing, re-centrifuging, adding the elution buffer, re-centrifuging and removing the elute. Sometimes a hybrid method is employed such that the binding is done by the batch method, but the solid phase with the target molecule bound is packed onto a column and washing and elution are done on the column. A third method, expanded bed absorption, which combines the advantages of the two methods mentioned above, has also been developed. The solid phase particles are placed in a column where liquid phase is pumped in from the bottom and exits at the top. The gravity of the particles ensure that the solid phase does not exit the column with the liquid phase. Affinity columns can be eluted by changing salt concentrations, pH, pI, charge and ionic strength directly or through a gradient to resolve the particles of interest. More recently, setups employing more than one column in series have been developed. The advantage compared to single column setups is that the resin material can be fully loaded, since non-binding product is directly passed on to a consecutive column with fresh column material. The resin costs per amount of produced product can thus be drastically reduced. Since one column can always be eluted and regenerated while the other column is loaded, already two columns are sufficient to make full use of the advantages. [1] Additional columns can give additional flexibility for elution and regeneration times, at the cost of additional equipment and resin costs.

Specific uses: Affinity chromatography can be used in a number of applications, including nucleic acid purification, protein purification from cell free extracts, and purification from blood.

Various affinity media exist for a variety of possible uses.[2] Briefly, they are (generalized): Activated/Functionalized – Works as a functional spacer, support matrix, and eliminates handling of

toxic reagents. Amino Acid – Used with a variety of serum proteins, proteins, peptides, and enzymes, as well as

rRNA and dsDNA. Avidin Biotin – Used in the purification process of biotin/avidin and their derivatives. Carbohydrate Bonding – Most often used with glycoproteins or any other carbohydrate-containing

substance. Carbohydrate – Used with lectins, glycoproteins, or any other carbohydrate metabolite protein. Dye Ligand – This media is nonspecific, but mimics biological substrates and proteins. Glutathione – Useful for separation of GST tagged recombinant proteins.

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Heparin – This media is a generalized affinity ligand, and it is most useful for separation of plasma coagulation proteins, along with nucleic acid enzymes and lipases.

Hydrophobic Interaction – Most commonly used to target free carboxyl groups and proteins. Immunoaffinity – Detailed below, this method utilizes antigens' and antibodies' high specificity to

separate. Immobilized Metal Affinity Chromatography – Detailed further below, this method uses interactions

between metal ions and proteins (usually specially tagged) to separate. Nucleotide/Coenzyme – Works to separate dehydrogenases, kinases, and transaminases. Nucleic Acid – Functions to trap mRNA, DNA, rRNA, and other nucleic acids/oligonucleotides. Protein A/G – This method is used to purify immunoglobulins. Speciality – Designed for a specific class or type of protein/coenzyme, this type of media will only

work to separate a specific protein or coenzyme.Immunoaffinity: Another use for the procedure is the affinity purification of antibodies from blood

serum. If serum is known to contain antibodies against a specific antigen (for example if the serum comes from an organism immunized against the antigen concerned) then it can be used for the affinity purification of that antigen. This is also known as Immunoaffinity Chromatography. Immobilized metal ion affinity chromatography: Immobilized metal ion affinity chromatography (IMAC) is based on the specific coordinate covalent bond of amino acids, particularly histidine, to metals. This technique works by allowing proteins with an affinity for metal ions to be retained in a column containing immobilized metal ions, such as cobalt, nickel, copper for the purification of histidine containing proteins or peptides, iron, zinc or gallium for the purification of phosphorylated proteins or peptides. Many naturally occurring proteins do not have an affinity for metal ions, therefore recombinant DNA technology can be used to introduce such a protein tag into the relevant gene. Methods used to elute the protein of interest include changing the pH, or adding a competitive molecule, such as imidazole.

Recombinant proteins: Possibly the most common use of affinity chromatography is for the purification of recombinant proteins. Proteins with a known affinity are protein tagged in order to aid their purification. The protein may have been genetically modified so as to allow it to be selected for affinity binding; this is known as a fusion protein. Tags include glutathione-S-transferase (GST), hexahistidine (His), and maltose binding protein (MBP). Histidine tags have an affinity for nickel or cobalt ions which have been immobilized by forming coordinate covalent bonds with a chelator incorporated in the stationary phase. For elution, an excess amount of a compound able to act as a metal ion ligand, such as imidazole, is used. GST has an affinity for glutathione which is commercially available immobilized as glutathione agarose. During elution, excess glutathione is used to displace the tagged protein.

Lectins: Lectin affinity chromatography is a form of affinity chromatography where lectins are used to separate components within the sample. Lectins, such as Concanavalin A[4] are proteins which can bind specific carbohydrate (sugar) molecules. The most common application is to separate glycoproteins from non-glycosylated proteins, or one glycoform from another glycoform.

PAPER CHROMATOGRAPHY: Paper chromatography is an analytical method used to separate colored chemicals or substances. It is primarily used as a teaching tool, having been replaced by other

chromatography methods, such as thin-layer chromatography. A paper chromatography variant, two-dimensional chromatography involves using two solvents and rotating the paper 90° in between. This is useful for separating complex mixtures of compounds having similar polarity, for example, amino acids. The setup has three components. The mobile phase is a solution that travels up the stationary phase, due to capillary action. The mobile phase is generally an alcohol solvent mixture, while the stationary phase is a strip of chromatography paper, also called a chromatogram.a chromatographic method is called adsorption chromatography if the stationary phase is solid.

Rƒ value, solutes, and solvents: The retardation factor (Rƒ) may be defined as the ratio of

the distance traveled by the substance to the distance traveled by the solvent. Rƒ values are usually

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expressed as a fraction of two decimal places. If Rƒvalue 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. To calculate the R ƒvalue, take the distance traveled by the substance divided by the distance traveled by the solvent (as mentioned earlier in terms of ratios). For example, if a compound travels 9.9 cm and the solvent front travels 12.7 cm, (9.9/12.7) the Rƒ value = 0.779 or 0.78. Rƒ value depends on temperature and the solvent used in experiment, so several solvents offer several Rƒ values for the same mixture of compound. A solvent in chromatography is the liquid the paper is placed in, and the solute is the ink which is being separated.

Types of Paper Chromatography:1. Descending Paper Chromatography-In this type, development of the chromatogram is done by allowing the solvent to travel down the paper. Here, mobile phase is placed in solvent holder at the top. The spot is kept at the top and above solvent flow down the paper from above.2. Ascending Paper Chromatography-Here the solvent travels up the chromatographic paper. Both Descending and Ascending Paper Chromatography are used for the separation of organic and inorganic substances.3. Ascending-Descending Paper Chromatography-It is the hybrid of both of the above techniques. The upper part of Ascending Chromatography can be folded over a rod in order to allow the paper to become Descending after crossing the rod.4. Radial Paper Chromatography-It is also called Circular Chromatography. Here a circular filter paper is taken and the sample is deposited at the center of the paper. After drying the spot, the filter paper is tied horizontally on a Petri dish containing solvent, so that the wick of the paper is dipped in the solvent. The solvent rises through the wick and the components are separated into concentric circles.5. Two-Dimensional Paper Chromatography-In this technique a square or rectangular paper is used. Here the sample is applied to one of the corners and development is performed at right angle to the direction of the first run.

Applications: Paper chromatography is one method for testing the purity of compounds and identifying substances. Paper chromatography is a useful technique because it is relatively quick and requires only small quantities of material. Separations in paper chromatography involve the same principles as those in thin layer chromatography, as it is a type of thin layer chromatography. In paper chromatography, substances are distributed between a stationary phase and a mobile phase. The stationary phase is the water trapped between the cellulose fibers of the paper. The mobile phase is a developing solution that travels up the stationary phase, carrying the samples with it. Components of the sample will separate readily according to how strongly they adsorb onto the stationary phase versus how readily they dissolve in the mobile phase. When a colored chemical sample is placed on a filter paper, the colors separate from the sample by placing one end of the paper in a solvent. The solvent diffuses up the paper, dissolving the various molecules in the sample according to the polarities of the molecules and the solvent. If the sample contains more than one color, that means it must have more than one kind of molecule. Because of the different chemical structures of each kind of molecule, the chances are very high that each molecule will have at least a slightly different polarity, giving each molecule a different solubility in the solvent. The unequal solubility causes the various color molecules to leave solution at different places as the solvent continues to move up the paper. The more soluble a molecule is, the higher it will migrate up the paper. If a chemical is very non-polar it will not dissolve at all in a very polar solvent. This is the same for a very polar chemical and a very non-polar solvent. It is very important to note that when using water (a very polar substance) as a solvent, the more polar the color, the higher it will rise on the paper.

THIN-LAYER CHROMATOGRAPHY: Thin-layer chromatography (TLC) is

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a chromatography technique used to separate non-volatile mixtures. Thin-layer chromatography is performed on a sheet of glass, plastic, or aluminium foil, which is coated with a thin layer of adsorbent material, usually silica gel, aluminium oxide (alumina), or cellulose. This layer of adsorbent is known as the stationary phase. After the sample has been applied on the plate, a solvent or solvent mixture (known as the mobile phase) is drawn up the plate via capillary action. Because different analytes ascend the TLC plate at different rates, separation is achieved. The mobile phase has different properties from the stationary phase. For example, with silica gel, a very polar substance, non-polar mobile phases such as heptane are used. The mobile phase may be a mixture, allowing chemists to fine-tune the bulk properties of the mobile phase. After the experiment, the spots are visualized. Often this can be done simply by projecting ultraviolet light onto the sheet; the sheets are treated with a phosphor, and dark spots appear on the sheet where compounds absorb the light impinging on a certain area. Chemical processes can also be used to visualize spots; anisaldehyde, for example, forms colored adducts with many compounds, and sulfuric acid will char most organic compounds, leaving a dark spot on the sheet. To quantify the results, the distance traveled by the substance being considered is divided by the total distance traveled by the mobile phase. (The mobile phase must not be allowed to reach the end of the stationary phase.) This ratio is called the retention factor or Rf. In general,a substance whose structure resembles the stationary phase will have low R f, while one that has a similar structure to the mobile phase will have high retention factor. Retention factors are characteristic, but will change depending on the exact condition of the mobile and stationary phase. For this reason, chemists usually apply a sample of a known compound to the sheet before running the experiment.

Separation Process and Principle: Different compounds in the sample mixture travel at different rates due to the differences in their attraction to the stationary phase and because of differences in solubility in the solvent. By changing the solvent, or perhaps using a mixture, the separation of components (measured by the Rf value) can be adjusted. Also, the separation achieved with a TLC plate can be used to estimate the separation of a flash chromatographycolumn. (A compound elutes from a column when the amount of solvent collected is equal to 1/R f.) Chemists often use TLC to develop a protocol for separation by chromatography and they use TLC to determine which fractions contain the desired compounds. Separation of compounds is based on the competition of the solute and the mobile phase for binding places on the stationary phase. For instance, if normal-phase silica gel is used as the stationary phase, it can be considered polar. Given two compounds that differ in polarity, the more polar compound has a stronger interaction with the silica and is, therefore, more capable to dispel the mobile phase from the binding places. As a consequence, the less polar compound moves higher up the plate (resulting in a higher Rf value).[6] If the mobile phase is changed to a more polar solvent or mixture of solvents, it is more capable of dispelling solutes from the silica binding places, and all compounds on the TLC plate will move higher up the plate. It is commonly said that "strong" solvents (eluents) push the analyzed compounds up the plate, whereas "weak" eluents barely move them. The order of strength/weakness depends on the coating (stationary phase) of the TLC plate. For silica gel-coated TLC plates, the eluent strength increases in the following order: perfluoroalkane (weakest), hexane, pentane, carbon tetrachloride, benzene/toluene, dichloromethane, diethyl ether, ethyl acetate, acetonitrile, acetone, 2 propanol/n butanol, water, methanol, triethylamine, acetic acid, formic acid (strongest). For C18-coated plates the order is reverse. In other words, when the stationary phase is polar and the mobile phase is polar, the method is normal-phase as opposed to reverse-phase. This means that if a mixture of ethyl acetate and hexane as the mobile phase is used, adding more ethyl acetate results in higher Rf values for all compounds on the TLC plate. Changing the polarity of the mobile phase will normally not result in reversed order of running of the compounds on the TLC plate. An eluotropic series can be used as a guide in selecting a mobile phase. If a reversed order of running of the compounds is desired, an apolar stationary phase should be used, such as C18-functionalized silica.

Applications: Characterization: In organic chemistry, reactions are qualitatively monitored with TLC. Spots sampled with a capillary tube are placed on the plate: a spot of starting material, a spot from the reaction mixture, and a cross-spot with both. This method is much more sensitive than the others and can be used to detect an extremely small amount of a compound, provided that it carries a radioactive atom. Isolation: The separated compounds each occupying a specific area on the plate, they can be scraped away, put in another solvent to separate them from the stationary phase and used for further analysis. Column Chromatography: Column chromatography is frequently used by organic chemists to purify liquids (and solids). An impure sample is loaded onto a column of adsorbent, such as silica gel or alumina. Usually, one starts with a less polar solvent to remove the less polar compounds, and then slowly

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increase the polarity of the solvent to remove the more polar compounds. Molecules with different polarity partition to different extents, and therefore move through the column at different rates. The eluent is collected in fractions.

GAS CHROMATOGRAPHY (GC): Gas chromatography is a common type of chromatography used in analytical chemistry for separating and analyzing compounds that can be vaporized without decomposition. Typical uses of GC include testing the purity of a particular substance, or separating the different components of a mixture (the relative amounts of such components can also be determined). In some situations, GC may help in identifying a compound. In preparative chromatography, GC can be used to prepare pure compounds from a mixture. In gas chromatography, the mobile phase (or "moving phase") is a carrier gas, usually an inert gas such as helium or an unreactive gas such as nitrogen. Helium remains the most commonly used carrier gas in about 90% of instruments although hydrogen is preferred for improved separations.[3] The stationary phase is a microscopic layer of liquid or polymer on an inert solid support, inside a piece

of glass or metal tubing called a column (an homage to the fractionating column used in distillation). The instrument used to perform gas chromatography is called a gas chromatograph (or "aerograph", "gas separator"). The gaseous compounds being analyzed interact with the walls of the column, which is coated with a stationary phase. This causes each compound to elute at a different time, known as the retention time of the compound. The comparison of retention times is what gives GC its analytical usefulness. Gas chromatography is in principle similar to column chromatography (as well as other forms of chromatography, such as HPLC, TLC), but has several notable differences. First, the process of separating the compounds in a mixture is carried out between a liquid stationary phase and a gas mobile phase, whereas in column chromatography the stationary phase is a solid and the mobile phase is a liquid. (Hence the full name of the procedure is "Gas–liquid chromatography", referring to the mobile and stationary phases, respectively.) Second, the column through which the gas phase passes is located in an oven where the temperature of the gas can be controlled, whereas column chromatography (typically) has no such temperature control. Finally, the concentration of a compound in the gas phase is solely a function of the vapor pressure of the gas. Gas chromatography is also similar to fractional distillation, since both processes separate the components of a mixture primarily based on boiling point (or vapor pressure) differences. However, fractional distillation is typically used to separate components of a mixture on a large scale, whereas GC can be used on a much smaller scale (i.e. microscale). Gas chromatography is also sometimes known as vapor-phase chromatography (VPC), or gas–liquid partition chromatography (GLPC). These alternative names, as well as their respective abbreviations, are frequently used in scientific literature. Strictly speaking, GLPC is the most correct terminology, and is thus preferred by many authors.

Method: This image above shows the interior of a Geo-Strata 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.

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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.

Applications: In general, substances that vaporize below 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 known as a reference standard. 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 analyse 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 a 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 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 two separate ones. The sensitivity is less, but this is a trade-off for simplicity in the gas supply. Gas Chromatography is used extensively in forensic science. Disciplines as diverse as solid drug dose (pre-consumption form) identification and quantification, arson investigation, paint chip analysis, and toxicology cases, employ GC to identify and quantify various biological specimens and crime-scene evidence.

HIGH PERFORMANCE LIQUID CHROMATOGRAPHY (HPLC): formerly referred as high-pressure liquid chromatography, is a technique in analytical chemistry used to separate, identify, and quantify each component in a mixture. It relies on pumps to pass a pressurized liquid solvent containing the sample mixture through a column filled with a solid adsorbent material. Each component in the sample interacts slightly differently with the adsorbent material, causing different flow rates for the different components and leading to the separation of the components as they flow out the column. HPLC has been used for manufacturing (e.g. during the production process of pharmaceutical and biological products), legal (e.g. detecting performance enhancement drugs in urine), research (e.g. separating the components of a complex biological sample, or of similar synthetic chemicals from each other), and medical (e.g. detecting vitamin D levels in blood serum) purposes.

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Chromatography can be described as a mass transfer process involving adsorption. HPLC relies on pumps to pass a pressurized liquid and a sample mixture through a column filled with adsorbent, leading to the separation of the sample components. The active component of the column, the adsorbent, is typically a granular material made of solid particles (e.g. silica, polymers, etc.), 2–50 micrometers in size. The components of the sample mixture are separated from each other due to their different degrees of interaction with the adsorbent particles. The pressurized liquid is typically a mixture of solvents (e.g. water, acetonitrile and/or methanol) and is referred to as a "mobile phase". Its composition and temperature play a major role in the separation process by influencing the interactions taking place between sample components and adsorbent. These interactions are physical in nature, such as hydrophobic (dispersive), dipole–dipole and ionic, most often a combination.

HPLC is distinguished from traditional ("low pressure") liquid chromatography because operational pressures are significantly higher (50–350 bar), while ordinary liquid chromatography typically relies on the force of gravity to pass the mobile phase through the column. Due to the small sample amount separated in analytical HPLC, typical column dimensions are 2.1–4.6 mm diameter, and 30–250 mm length. Also HPLC columns are made with smaller sorbent particles (2–50 micrometer in average particle size). This gives HPLC superior resolving power (the ability to distinguish between compounds) when separating mixtures, which makes it a popular chromatographic technique. The schematic of an HPLC instrument typically includes a sampler, pumps, and a detector. The sampler brings the sample mixture into the mobile phase stream which carries it into the column. The pumps deliver the desired flow and composition of the mobile phase through the column. The detector generates a signal proportional to the amount of sample component emerging from the column, hence allowing for quantitative analysis of the sample components. A digital microprocessor and user software control the HPLC instrument and provide data analysis. Some models of mechanical pumps in a HPLC instrument can mix multiple solvents together in ratios changing in time, generating a composition gradient in the mobile phase. Various detectors are in common use, such as UV/Vis, photodiode array (PDA) or based on mass spectrometry. Most HPLC instruments also have a column oven that allows for adjusting the temperature at which the separation is performed

Types: Partition chromatography: Partition chromatography was one of the first kinds of chromatography that chemists developed. The partition coefficient principle has been applied in paper chromatography, thin layer chromatography, gas phase and liquid–liquid separation applications. Use of more polar solvents in the mobile phase will decrease the retention time of the analytes, whereas more hydrophobic solvents tend to increase retention times. Normal–phase chromatography: Also known as normal-phase HPLC (NP-HPLC) this method separates analytes based on their affinity for a polar stationary surface such as silica, hence it is based on analyte ability to engage in polar interactions (such as hydrogen-bonding or dipole-dipole type of interactions) with the sorbent surface. Displacement chromatography: A molecule with a high affinity for the chromatography matrix (the displacer) will compete effectively for binding sites, and thus displace all molecules with lesser affinities. Reversed-phase chromatography (RPC): It has a non-polar stationary phase and an aqueous, moderately polar mobile phase. One common stationary phase is a silica which has been surface-modified with RMe 2SiCl, where R is a straight chain alkyl group such as C18H37 or C8H17. Size-exclusion chromatography: Is also known as gel permeation chromatography or gel filtration chromatography, separates particles on the basis of molecular size (actually by a particle's Stokes radius). It is generally a low resolution chromatography and thus it is often reserved for the final, "polishing" step of the purification. Ion-exchange chromatography: is based on the attraction between solute ions and charged sites bound to the stationary phase. Solute ions of the same charge as the charged sites on the column are excluded from binding, while solute ions of the opposite charge of the charged sites of the column are retained on the column.

Applications: Manufacturing: As briefly mentioned, HPLC has many applications in both laboratory and clinical science. It is a common technique used in pharmaceutical development as it is a dependable way to obtain and ensure product purity. Legal: This technique is also used for detection of illicit drugs in urine. The most common method of drug detection is an immunoassay. Research: detecting concentrations of potential clinical candidates like anti-fungal and asthma drugs. Medical: While urine is the most common medium for analyzing drug concentrations, blood serum is the sample collected for most medical analyses with HPLC.

ELECTROPHORESIS:

PRINCIPLE: The surface adsorbed sample strongly affects suspended particles by applying

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electric surface charge, on which an external electric field exerts an electrostatic coulomb force. According to the double layer theory, all surface charges in fluids are screened by a diffuse layer of ions, which has the same absolute charge but opposite sign with respect to that of the surface charge. The electric field also exerts a force on the ions in the diffuse layer which has direction opposite to that acting on the surface charge This force is not actually applied to the particle, but to the ions in the diffuse layer located at some distance from the particle surface, and part of it is transferred all the way to the particle surface through viscous stress. This part of the force is also called electrophoretic retardation force. When the electric field is applied and the charged particle to be analyzed is at steady movement through the diffuse layer, the total resulting force is zero. Considering the drag force on the moving particles due to the viscosity of the dispersant, in the case of low turbulence and moderate electric charge strength E, the velocity of a dispersed particle v is simply proportional to the applied field, which leaves the electrophoretic mobility |i e

defined as. The most known and widely used theory of electrophoresis was developed in 1903 by Smoluchowsky. where er is the dielectric constant of the dispersion, e0 is the permittivity of free space (C2

N-11 m-2), n is dynamic viscosity of the dispersion medium (Pa s), and Z is zeta potential (i.e., the electrokinetic potential of the slipping plane in the double layer). The Smoluchowski theory is very powerful because it works for dispersed particles of any shape at any concentration. Unfortunately, it has limitations on its validity. It follows, for instance, from the fact that it does not include Debye length k-1. However, Debye length must be important for electrophoresis, as follows immediately from the Figure on the right. Increasing thickness of the double layer (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 the Smoluchowski theory is valid only for sufficiently thin DL, when particle radius a is much greater than the Debye length. 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. The Smoluchowski theory also neglects contribution of surface conductivity. This is expressed in modern theory as condition of small Dukhin number. In the effort of expanding the range of validity of electrophoretic theories, the opposite asymptotic case was considered, when Debye length is larger than particle radius: Under this condition of a “thick Double Layer”, Huckel predicted the following relation for electrophoretic mobility: This model can be useful for some nanoparticles and non-polar fluids, where Debye length is much larger than in the usual cases. There are several analytical theories that incorporate surface conductivity and eliminate the restriction of a small Dukhin number pioneered by Overbeek and Booth. Modern, rigorous theories valid for any zeta potential and often any aK stem mostly from Dukhin- Semenikhin theory. In the thin Double Layer limit, these theories confirm the numerical solution to the problem provided by O’Brien and White.

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POLYACRYLAMIDE GEL ELECTROPHORESIS: (PAGE) describes a technique widely used in biochemistry, forensics, genetics, molecular biology and biotechnology to separate biological macromolecules, usually proteins or nucleic acids, according to their electrophoretic mobility. Mobility is a function of the length, conformation and charge of the molecule. As with all forms of gel electrophoresis, molecules may be run in their native state, preserving the molecules' higher-order structure, or a chemical denaturant may be added to remove this structure and turn the molecule into an unstructured linear chain whose mobility depends only on its length and mass-to-charge ratio. For nucleic acids, urea is the most commonly used denaturant. SDS-PAGE (Sodium Dodecyl Sulfate - Polyacrylamide Gel Electrophoresis): One of the most common means of analyzing proteins by electrophoresis is by using Sodium Dodecyl Sulfate - Polyacrylamide Gel Electrophoresis. SDS is a detergent which denatures proteins by binding to the hydrophobic regions and essentially coating the linear protein sequence with a set of SDS molecules. The SDS is negatively charged and thus becomes the dominant charge of the complex. The number of SDS molecules that bind is simply proportional to the size of the protein. Therefore the charge to mass ratio should not change with size. DNA Agarose Gels…A simple way of separating fairly large fragments of DNA from one another by size is to use an agarose gel. DNA denaturing polyacrylamide gels (often called sequencing gels)… To look at smaller DNA molecules with much higher resolution, people generally denature the DNA via heat and run it through a thin polyacrylamide gel that is also kept near the denaturing temperature. These gels usually contain additional denaturing compounds such as Urea. Two pieces of DNA that differ in size by 1 base can be distinguished from each other this way.

Polyacrylamide gel (PAG) had been known as a potential embedding medium for sectioning tissues as early as 1964, and two independent groups employed PAG in electrophoresis in 1959. It possesses several electrophoretically desirable features that make it a versatile medium. It is a synthetic, thermo-stable, transparent, strong, chemically relatively inert gel, and can be prepared with a wide range of average pore sizes.[13] The pore size of a gel is determined by two factors, the total amount of acrylamide present (%T) (T = Total concentration of acrylamide and bisacrylamide monomer) and the amount of cross-linker (%C) (C = bisacrylamide concentration). Pore size decreases with increasing %T; with cross-linking, 5%C gives the smallest pore size. Any increase or decrease in %C from 5% increases the pore size, as pore size with respect to %C is a parabolic function with vertex as 5%C. This appears to be because of non-homogeneous bundling of polymer strands within the gel. This gel material can also withstand high voltagegradients, is amenable to various staining and destaining procedures, and can be digested to extract separated fractions or dried for autoradiography and permanent recording.

Applications: Various buffer systems are used in PAGE depending on the nature of the sample and the experimental objective. The buffers used at the anode and cathode may be the same or different . An electric field is applied across the gel, causing the negatively charged proteins or nucleic acids to migrate across the gel away from the negative electrode (which is the cathode being that this is an electrolytic rather than galvanic cell) and towards the positive electrode (the anode). Depending on their size, each biomolecule moves differently through the gel matrix: small molecules more easily fit through the pores in the gel, while larger ones have more difficulty. The gel is run usually for a few hours, though this depends on the voltage applied across the gel; migration occurs more quickly at higher voltages, but these results are typically less accurate than at those at lower voltages. After the set amount of time, the biomolecules have migrated different distances based on their size. Smaller biomolecules travel farther down the gel, while larger ones remain closer to the point of origin. Biomolecules may therefore be separated roughly according to size, which depends mainly on molecular weight under denaturing conditions, but also depends on higher-order conformation under native conditions. However, certain glycoproteins behave anomalously on SDS gels.

AGAROSE GEL ELECTROPHORESIS: is a method of gel electrophoresis used in biochemistry, molecular biology, genetics, and clinical chemistry to separate a mixed population of macromolecules such as DNA or proteins in a matrix of agarose, one of the two main components of agar. The proteins may be separated by charge and/or size (isoelectric focusing agarose electrophoresis is essentially size independent), and the DNA and RNA fragments by length. Biomolecules are separated by applying an electric field to move the charged molecules through an agarose matrix, and the biomolecules are

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separated by size in the agarose gel matrix. Agarose gel is easy to cast, has relatively fewer charged groups, and is particularly suitable for separating DNA of size range most often encountered in laboratories, which accounts for the popularity of its use. The separated DNA may be viewed with stain, most commonly under UV light, and the DNA fragments can be extracted from the gel with relative ease. Most agarose gels used are between 0.7 - 2% dissolved in a suitable electrophoresis buffer.

Mechanism of migration and separation: The negative charge of its phosphate backbone moves the DNA towards the positively charged anode during electrophoresis. However, the migration of DNA molecules in solution, in the absence of a gel matrix, is independent of molecular weight during electrophoresis. The gel matrix is therefore responsible for the separation of DNA by size during electrophoresis, and a number of models exist to explain the mechanism of separation of biomolecules in gel matrix. A widely accepted one is the Ogston model which treats the polymer matrix as a sieve. A globular protein or a random coil DNA moves through the interconnected pores, and the movement of larger molecules is more likely to be impeded and slowed down by collisions with the gel matrix, and the molecules of different sizes can therefore be separated in this sieving process. The Ogston model however breaks down for large molecules whereby the pores are significantly smaller than size of the molecule. For DNA molecules of size greater than 1 kb, a reptation model (or its variants) is most commonly used. This model assumes that the DNA can crawl in a "snake-like" fashion (hence "reptation") through the pores as an elongated molecule. A biased reptation model applies at higher electric field strength, whereby the leading end of the molecule become strongly biased in the forward direction and pulls the rest of the molecule along. Real-time fluorescence microscopy of stained molecules, however, showed subtler dynamics during electrophoresis, with the DNA showing considerable elasticity as it alternately stretching in the direction of the applied field and then contracting into a ball, or becoming hooked into a U-shape when it gets caught on the polymer fibres.

Requirements…..Electrophoretic unit, conical flask, measuring cylinder, power pack, micropipette, micro tips (1X) TAE buffer, Gel loading dye, EtBr, Agarose. Steps…..For preparing 0.8% agarose gel, 0.14g of Agarose was dissolved in 20ml of TAE (1X). Mixture was boiled till a clear solution was obtained. Left at room temperature till suspension reaches 40-45°C. Then 2pl of 1% EtBr was added. Seal the casting tray properly and placed the combs at appropriate place. Pour the gel and leave at room temperature for 45-50 minutes to solidify the gel. Fill the buffer tank with TAE (1X) so that the gel was dipped. 5 pl of the sample was loaded into the well by mixing with 1pl of 6X loading dye containing Bromophenol blue. Switch on the power supply at the rate of 5V/cm. When the electrophoretic front reaches bottom of the gel power supply was switched off. The gel was placed over transilluminator and observed under UV light.

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 DNA fragments for extraction and purification. Separation of restricted genomic DNA prior to Southern transfer, or of RNA prior to Northern transfer. Agarose gels are easily cast and handled compared to other matrices and nucleic acids are not chemically altered during electrophoresis. Samples are also easily recovered. After the experiment is finished, the resulting gel can be stored in a plastic bag in a refrigerator. Electrophoresis is performed in buffer solutions to reduce pH changes due to the electric field, which is important because the charge of DNA and RNA depends on pH, but running for too long can exhaust the buffering capacity of the solution. Further, different preparations of genetic material may not migrate consistently with each other, for morphological or other reasons. Agarose Gel electrophoresis is used in forensics, molecular biology, genetics, microbiology and biochemistry. The results can be analyzed quantitatively by visualizing the gel with UV light and a gel imaging device. The image is recorded with a computer operated camera, and the intensity of the band or spot of interest is measured and compared against standard or markers loaded on the same gel. Depending on the type of analysis being performed, other techniques are often implemented in conjunction with the results of gel electrophoresis, providing a wide range of field-specific applications. Electrophoresis plays a vital role in the separation of nucleic acids and proteins in the field of genomics and proteomics. The techniques are very simple but has its role in advanced studies. The electrophoretic devices are economical and can be explored to the core to analyze the complexity of biomolecules. The movement of particles under spatially uniform electric field in a fluid is called electrophoresis. Polyacrylamide and agarose… It is used to visualize the Et Br stained DNA or RNA in gel through ultraviolet light of specific wavelength. Bromophenol blue and xylene cyanol, TAE (tris Acetate EDTA) or TBE (Tris Borate EDTA) SDS - Sodium dodecyl

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sulphate, PAGE - polyacrylamide gel electrophoresis, CE - capillary electrophoresis, Sequencing gels, Gel has a pH gradient, Separate large DNA fragments, ,Separate protein without SDS, Separate proteins based on electrophoresis and reaction with antibodies, separate ionic species by charge, friction force and hydrodynamic radius.

ELECTROFOCUSING also known as ISOELECTRIC FOCUSING (IEF): is a technique for separating different molecules by differences in their isoelectric point (pI). It is a type of zone electrophoresis, usually performed on proteins in a gel, that takes advantage of the fact that overall charge on the molecule of interest is a function of the pH of its surroundings. Isoelectric focusing (IEF) is an electrophoretic technique for the separation of proteins based on their isoelectric point (pI). The pI is the pH at which a protein has no net charge and thus, does not migrate further in an electric field. IEF gels are used to determine the pI of a protein and to detect minor changes in the protein due to post-translational modifications such as phosphorylation and glycosylation.

How isoelectric focusing (IEF) works: SDS-PAGE is the standard technique used for separation of proteins in the lab, but that doesn’t meant that other techniques don’t have their place–one such technique is isoelectric focusing (IEF). IEF, also known simply as electrofocusing, is a technique for separating charged molecules, usually proteins or peptides, on the basis of their isoelectric point (pI), i.e., the pH at which the molecule has no charge. IEF works because in an electric field molecules in a pH gradient will migrate towards their pI. In most cases, a commercially available immobilized pH gradient (IPG) strip is used. The IPG strip consists of an acrylamide gel that contains wide pores to prevent a sieving effect based on protein mass, with a pH gradient. Various gradients are available, with wider gradients, such as pH 3-10 that are used for whole proteome analysis, and narrower ranges, such as pH 5-8 that are used for more specialist applications. The sample is usually combined with carrier ampholytes to assist in migration. Ampholytes are a mixture of charged molecules with a range of pIs that matches the pI range of the IPG strip. The migration of the ampholytes encourages the sample molecules to move along the pH gradient. Ampholyte mixtures of a variety of pI ranges are commercially available. After separation across the pH gradient, the sample is further separated (in 2D-PAGE) or analyzed (in the case of fractionation for mass spec–more on this below).

Applications of IEF: IEF is “traditionally” used as first stage separation for 2D-PAGE, separating proteins by charge prior to second dimension separation by SDS-PAGE. This additional separation allows resolution of a couple of thousand proteins on a 2D-PAGE gel–enough for the entire proteome of an organelle or bacterium. It can also be used to examine post-translation modifications of proteins. Another use for IEF is for fractionation of proteins or peptides prior to mass spec. Previously, it was difficult to recover molecules separated by IEF. However, with the Agilent OFFGEL system, the proteins or peptides remain in solution rather than being trapped in the gel as with standard IEF. Electrofocusing of proteins in this way provides a convenient alternative to SDS-PAGE for sample fractionation prior to mass spec. Using IEF to separate peptides also provides an alternative to strong cation exchange (SCX) fractionation, which is one of the most popular techniques for the separation of peptides prior to mass spectrometry. IEF separation of peptides has been shown to result in more peptide identifications from whole proteome samples than with SCX for some samples In isoelectric focusing (IEF), proteins are applied to polyacrylamide gels (IEF gels) or immobilized pH gradient (IPG) strips containing a fixed pH gradient. An electrical field is applied and the protein sample containing a mixture of proteins migrates through the pH gradient. Individual proteins are immobilized in the pH gradient as they approach their specific pI. After staining the gel and documenting the results, proteins separated by pI can be separated by mass using 2D gel electrophoresis.

ULTRACENTRIFUGATION:

An important tool in biochemical research is the centrifuge, which through rapid spinning imposes high centrifugal forces on suspended particles, or even molecules in solution, and causes separations of such matter on the basis of differences in weight. The ultracentrifuge is a centrifuge optimized for spinning a rotor at very high speeds, capable of generating acceleration as high as 1 000 000 g (approx. 9 800 km/s²).[1]There are two kinds of ultracentrifuges, the preparative and the analytical ultracentrifuge. Both classes of instruments find important uses in molecular biology, biochemistry, and polymer science. Example: Red cells may be separated from plasma of blood, nuclei from mitochondria in cell homogenates,

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and one protein from another in complex mixtures. It is the apparent outward force that draws a rotating body away from the centre of rotation. It is caused by the inertia of the body as the body's path is continually redirected. (Inertia - it is the resistance of any physical object to a change in its state of motion or rest, or the tendency of an object to resist any change in its motion.). Analytical Ultracentrifuge: In an analytical ultracentrifuge, a sample being spun can be monitored in real time through an optical detection system, using ultraviolet light absorption and optical refractive index sensitive system. As the rotor turns, the images of the cell (proteins) are projected by an optical system on to film or a computer. The concentration of the solution at various points in the cell is determined by absorption of a light of the appropriate wavelength. Reparative ultracentrifuges: Swinging bucket rotors allow the tubes to hang on hinges so the tubes reorient to the horizontal as the rotor initially accelerates. Zonal rotors are designed to contain a large volume of sample in a single central cavity rather than in tubes. Some zonal rotors are capable of dynamic loading and unloading of samples while the rotor is spinning at high. 1) Differential ultracentrifugation: is a common procedure in microbiology and cytology used to separate certain organelles from whole cells for further analysis of specific parts of cells. 2) Density gradient: two types of density gradient centrifugations under preparative centrifugation such as: ZONAL (or) RATE, ISOPYCNIC, Mixture to be separated is layered on top of a SUCROSE, or FICOLL, GRADIENT (increasing concentration down the tube) provides gravitational stability as different species move down tube at different rates. Forming separate bands. Sedimenting force on particle = Mass x centrifugal field = mɯ2r. Species are separated by differences in SEDIMENTATION COEFFICIENT (S)=Rate of movement down tube/ Centrifugal force is increased for particle of LARGER MASS is also increased for MORE COMPACT STRUCTURES of equal particle mass. Mild, non-denaturing procedure, useful for protein purification, and for intact cells and organelles, Molecules separated on EQUILIBRIUM POSITION, NOT by RATES of sedimentation. Each molecule floats or sinks to position where density equals density of CsCl solution. Then no net sedimenting force on molecules. Isopycnic = Equal density and separation is on basis of DIFFERENT DENSITIES of the particles and usually in reference to a process of separating particles, sub cellular organelles.

Key Concepts: Centrifugation, in particular high‐speed (ultra) centrifugation, is a widely used technique to elucidate fundamental processes such as adenosine triphosphate (ATP) synthesis in mitochondria, the synthesis of proteins by ribosomes or the interactions of intracellular multiprotein complexes. Ultracentrifugation is basically carried out in two ways: preparative and analytical centrifugation. The former aims to isolate and purify, for example, subcellular organelles or multiprotein complexes; the latter allows to analyse interactions between macromolecules and to unravel physico‐chemical properties like mass and size of such particles. Preparative centrifugation performed as batch‐type (conventional) centrifugation is mostly used to separate and enrich organelles out of complex biological mixtures. The alternative, continuous‐flow centrifugation, is particularly useful for the large‐scale collection of particles out of a diluted solution as it combines high centrifugal force with high throughput. Theoretically, two kinds of preparative centrifugation have to be distinguished: differential

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centrifugation and density gradient centrifugation. Differential centrifugation fractionates organelles of a tissue homogenate according to their size and shape, yet leads only to an enriched rather than a highly purified preparation of a particular organelle. To get a preparation genuinely purified, common contaminants have to be subsequently removed by density gradient centrifugation. Rate zonal and isopycnic density gradient centrifugations differ in their basic concepts and the types of density gradients used.

Ultracentrifuge applications: Ultracentrifuges are commonly used in molecular biology, biochemistry, and cell biology. Applications of ultracentrifuges include the separation of small particles such as viruses, viral particles, proteins and/or protein complexes, lipoproteins, RNA, and plasmid DNA. 3D Cell Culture, Apoptosis, Cell-Based Assays, Cell Maintenance, Cell Signaling, Dendritic Cells, Exosomes, High Throughput Screening, Immune Monitoring, Nanoparticle Research, Next Generation Sequencing, Sample Prep for Mass Spectrometry, Stem Cells. determine sample purity, characterize assembly and disassembly mechanisms of biomolecular complexes, determine subunit stoichiometries, detect and characterize macromolecular conformational changes, measure equilibrium constants and thermodynamic parameters for self- and hetero-associating systems, characterize the solution-state behavior of macromolecules under various conditions.

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