（ 1 ） Before entering the laboratory, you must make careful preview of the experiment to know the principles and the main steps. Those who fail to preview, will not be allowed to start the experiment until he or she makes it up.
（ 2 ） Do not eat food, drink beverages, or chew gum in the laboratory. Do not use laboratory glassware as containers for food or beverages. Never use mouth suction to fill a pipette. Use a pipette pump.
（ 3 ） Perform only those experiments authorized by the instructor. Follow all written and verbal instructions carefully. If you do not understand a direction or part of a procedure, ask the instructor before proceeding. Unauthorized experiments are prohibited.
（ 4 ） During the lab course, work areas should be kept clean and tidy at all times. you should keep quiet, operate regularly, observe carefully, think earnestly, strive for getting accurate and good results.
（ 5 ） Record the observation and data timely and directly on your notebook. You must turn in your notebook on time.
（ 6 ） All reagent bottles and instruments should be returned to the primary situs immediately after use. All glass wares should be washed clean before and after use.
（ 7 ） Do not use the sink to discard matches, filter paper or insoluble solids. Use the waste containers that are provided.
（ 8 ） Labels and equipment instructions must be read carefully before use. Set up and use the prescribed apparatus as directed in the laboratory instructions or by your instructor.
（ 9 ） The students on duty should clean the lab, set things in order and check the safety of water taps, electricity, windows and doors before leaving the lab.
（ 1 ） Report any accident (spill, breakage, etc.) or injury (cut, burn, etc.) to the instructor immediately, no matter how trivial it may appear.
（ 2 ） If a chemical should splash in your eye(s) or on your skin, immediately flush with running water for at least 20 minutes.
（ 3 ） If there is a fire during a laboratory period; gas valves must be turned off, fume hoods turned off, and any electrical equipment turned off. Notify the instructor immediately.

（ 4 ） When mercury thermometers are broken, mercury must not be touched. Notify the instructor immediately.
（ 1 ） The experimental title
（ 2 ） A brief statement of the purpose
（ 3 ） A brief statement of the theory and principal manipulative steps
（ 4 ） The raw data
（ 5 ） Complete calculation
（ 6 ） Results and interpretations/discusion
（ 7 ） Conclusions if possible and necessary
(1)The use of pipette
Pipettes are used by research scientists to transfer small volumes of liquid from one vessel
(such as a test tube) to another. Pipette volumes can range from as small as 0.1 microliter up to 20 milliliters. Be aware that there are different graduations on different sizes of pipette. Make sure you use the correct size!
General use of pipette
 Hold the pipette with your thumb and middle finger. Squeeze the bulb with your left hand to create a vacuum.
 Put the tip of the pipette into the liquid, and the bulb upside the pipette . Release your grip on the bulb. Watch carefully, the liquid rises quickly. Fill the pipette to a level just higher than you need for the experiment. Then press upside of the pipette with forefinger tightly.
 Adjust the liquid level to the area you want to dispense the liquid. Move your pipette to the next container and let your forefinger go to release the fluid.
 Dispose of pipette properly when finished.
(2)Cleaning of Pipette
Not only can a dirty pipette contaminate your sample, it can also dispense the wrong volume!
Liquid sticks to the walls of a dirty pipette, so one test for a clean pipette is to check that the solution does not bead on the walls as it is dispensed. Unless you are being provided a new pipette by the technician, it is a good idea to wash your pipette as outlined in the following steps:
（ 1 ） Place a small volume of the washing solution into a beaker.
（ 2 ） Draw the solution into the pipette using a pipette pump, and then tilt and turn the pipette until all of the inner surfaces have been wetted by the solution.
（ 3 ） Discard this washing solution as waste.
（ 4 ） Repeat the previous steps again until beading is not observed on the inner walls of the pipette.

A micropipette is a precision pump fitted with a disposable tip. Almost all measurements of 1 ml (1000 µL ) or less are done using a micropipette. These are precision instruments costing more than $250 a piece, so be careful when using one.
(1)The kind of micropipette:
There are different size micropipettes with various volume range: a small-volume (1-20 ul), a mid-range (20-100 ul), and a large-volume (100-1000 ul). Depending on the volume needed, appropriate size micropipette should be used to ascertain accuracy. Since these micropipettes are very delicate, hard to repair and very costly, please note the following rules in using a micropipette.

Never rotate the volume adjustor beyond the upper or lower range of the pipette as stated by the manufacturer.

Never use the micropipettes without the tip in place; this could ruin the piston.

Never invert or lay the micopipette down with a filled tip; fluid could run back into the piston.

Never let the plunger snap back after withdrawing or expelling fluid. This could damage the piston.

Never reuse a tip that has been used to measure a different reagent.
1. Select the correct micropipette for the volume to be delivered. The range of the pipette can be found on the top of the pipette plunger. Rotate the volume adjustor to the desired setting.
2. Firmly seat a proper-sized tip on the end of the micropipette.
3. When withdrawing or expelling fluid, always hold the tube firmly between your thumb and forefinger. For best control, grasp the micropipette in your palm and wrap your fingers around the barrel; work the plunger with your thumb.
4. The micropipette plunger has two positions. If you want to withdraw fluid, depress the plunger to the first stop and hold it in this position. Then dip the tip into the solution to the approximate correct depth and hold the pipette perpendicular to the surface of the solution. Draw fluid into the tip by gradually releasing the plunger. When the plunger has completed its return, wait 1 additional second. Be sure the tip remains in the solution while you are releasing the plunger.
5. Slide the pipette tip along the inside wall of the reagent tube to dislodge any excess droplets adhering to the outside of the tip. Check and make sure there is no air space at the very tip of the tip.
6. Deliver the solution by placing the tip against the inside wall of the receiving container as close as possible to the liquid surface or just at the bottom if the container is empty. Slowly depress the plunger to the first stop pause 1~2seconds and continue to depress to the second stop to blow out the last bit of fluid. Hold the plunger in the depressed position and slide the tip out of the pipette along the inside wall of the reaction tube. This creates a capillary effect that helps draw the last bit of fluid out of the tip.
7. Eject the tip into the proper waste container by depressing the tip-ejection button.
8. Repeat the process using a fresh tip for each delivery.
Spectrophotometric analysis is the most widely used method of quantitative and qualitative analysis in the chemical and biological sciences; it is an accurate and very sensitive method which

can analyze quantities as small as micrograms. The method depends on the light absorbing properties of either the substance being analyzed or a derivative of its. The basis of spectrophotometry is simple: the intensity of the light which is transmitted through a solution containing an absorbing substance (chromogen) is decreased by that fraction which is absorbed, and this fraction can be detected and measured photoelectrically.
Visible light represents a very narrow section of this range of wavelengths between 400 nanometers (nm) for purple light to around 750 nm for red light. Shorter wavelengths fall into the ultraviolet region and longer wavelengths are in the infrared region. White light is a mixture of all of the wavelengths in the visible range. When light strikes an object, it may be reflected, absorbed, transmitted, or diffracted. A prism or a diffraction grating separates white light into its various colors. If some of the light is absorbed, the reflected or transmitted light has the complementary color of the absorbed light. A spectrophotometer uses an arrangement of prisms, mirrors, and slits to select light of a desired wavelength and to direct it toward a sample compartment and a detector.
The detector electronically measures the intensity of the light striking it. A sample is placed in the light path, and the instrument compares the intensity of the light going through the sample (I) to the intensity observed without the sample (I o
). A spectrophotometer is often used to study solutions. A solution containing an absorbing material is compared to a reference solution of the same solvent and non-absorbing materials. The transmittance of the reference solution is set to
100% (A = 0), then the relative transmittance or A value of the solution is measured.
Absorbtion may occur when light of the energy (wavelength) required to excite certain of the bonding electrons strikes the absorbing molecule. The intensity and position in the electromagnetic spectrum of the absorbed light is related in a complicated way to the electronic structure of the molecule and its constituent atoms and is a unique optical characteristic of the substance. A plot of the absorption of a solution of an absorbing substance over a range of wavelengths is referred to as the absorption spectrum of the compound and is useful for the qualitative identification of unknown compounds.
Spectrophotometry involves the qualitative and quantitative use of absorption data obtained from compounds that absorb light in the ultraviolet(UV<400nm), visible, and near infrared(>750nm) regions. Much of photometry is based on two formalized laws(Lambert’s law and Beer’s law).
If a beam of monochromatic light passed through a solution, part of the light is absorbed by the solution. The intensity of transmitted light is weaker than that of the incident light. When the concentration of the solution is fixed, the thicker the light path is, the weaker the intensity of transmitted light will be.

Let I
0
= intensity of the incident light, I = intensity of transmitted light, L = length of light path(thickness of the solution), then
I/I
0
= 10 -K
1
L or lgI/I
0
= K
1
L (1)
K
1
is the extinction coefficiet which is influenced by the wavelength of light, the nature of the solvent and the concentration of the solute in the solution. As shown in equation (1), the diminution of the intensity of incident light (I/I
0
), as the light passed through a solution, is not simply in proportion to the thickness of the solution, but has the relationship of exponential function with the thickness of the solution. This is the Lambert’s law.
If a beam of monochromatic light is passed through a solution with fixed thickness, the higher the concentration, the weaker the intensity of the transmitted light. The relationship between those is similar to Lambert’s law which is shown as follows:
I/I
0
= 10 -K
2
C or lgI/I
0
= K
2
C (2)
Where C = concentration of solution, K
2
is a constant influenced by the wavelength of light, the nature of solvent and the thickness of the solution. Equation (2) which expresses the relationship between light intensity and the concentration of solution is termed as Beer’s law.
Lambert’t and Beer’s law can be combined together as follows:
A = - lgI/I
0
= -lgT = KCL (3) where I and I
0
are the intensity of transmitted light in the presence and in the absence of the chromogen, respectively; C is the concentration of the chromogen; L is the length of the light path through the solution; K is a constant, characteristic for each absorbing substance at a specific wavelength of light and in a specified solvent. It should be remembered that a K value is given for a particular wavelength and solvent and that another K value applies at a different wavelength or with another solvent. The ratio I/I
0
is called the light transmittance(T) and is usually measured in percent. The absorbance(A), or optical density (O.D.), expresses the extent of absorption by solution. it is the quantity more frequently used.
The Beer-Lambert law states that the amount of light absorbed(A value) is proportional both to the concentration of the chromogen in solution and to the length of the light path through the solution.
In this form theLambert- Beer’s law states that doubling of either the concentration of the absorbing substance or doubling of the depth of solution leads to a doubling of the A. The quantitative measurement of the amount of light absorbed is accomplished with an instrument called a spectrophotometer. Spectrophotometers detect photoelectrically and compare electronically the difference in the amount of light transmitted through solutions containing differing concentrations of an absorbing substance. The difference is expressed on a dial of the instrument as percent transmission or absorbance.

(1) Standard tube method:
As known from equation (3), if solvent and wavelength are fixed(extinction coefficient is fixed), the absorbance of a solution is proportional to its concentration and the length of the light path.
A
1
/A
2
= C
1
/C
2
or C
2
= A
2
/A
1
*C
1
(4)
Where C
2
= concentration of sample solution, C
1
= concentration of standard solution, A
2
= absorbance of sample solution, A
1
= absorbance of standard solution. So C
2
can be calculated after measuring the absorbances of standard and sample solutions respectively.
(2) Standard curve method:
The quantitative measurement of an absorbing substance can also be accomplished using such a plot, called a standard curve. The curve is constructed by preparing a series of standards of the substance to be analyzed in a graded series of known concentrations. The A value of each standard is then determined by spectrophotometer. Plot a standard curve with the known concentrations of protein as the abscissa, and the corresponding absorptions as the ordinate. The A value of a solution of unknown concentration is then determined under identical conditions and the value of the concentration simply read from the plot. Two things must be understood about standard curves. First, they are called standard curves because the response of absorbance to the amount of added protein is normally not exactly linear or may be quite different from linear.
Second, the range of protein you include in your standard curve (e.g. 0 to 25 µg or 0 to 100 µg) will depend on the assay being used. You need to make sure that the unknown you are determining falls within your standard curve. It is usually desirable to have at least two points on either side of the determination of your unknown, and the more points on either side of your determination, the more accurate your results will be.
In practice Absorbance measurements are made by comparing a solution containing the sample to a reference solution. It is possible to define various references, but usually the reference is a blank, i.e., a solution identical in composition with the sample except that the absorbing material being measured is absent. In making this spectrophotometric measurement we set the instrument at 100 percent transmission (zero A) using the reference solution. The sample containing the chromogen to be measured is then placed in the light path and its A value relative to the reference solution is read.
You need to make sure that both your standard curve and your unknown determination fall within the absorbance limits of your spectrophotometer. Most spectrophotometers cannot linearly read absorbance values beyond 2.0 units. Thus, you may have to dilute your protein of interest to work within this range.
Ⅱ
1. Light source
The light source must be capable of emitting a steady amount of sufficient energy in the wavelength ranges required for analysis of the sample. Most photometers employ tungsten lamp for spectral analysis in the range of 400~900nm. But the photometers which have capability for

analysis in the UV range employ hydrogen lamp that emits light in the ranges of 200~400nm.
2. Monochromators
Monochromators are employed to generate the desired wavelength of light.
3. Cuvette
Sample cuvette is one of the most important components. Optical glass cuvettes are employed in visual range(400~700nm), and quartz cuvette in UV range(200~400nm). In order to keep the cuvette at very good working conditions, the optical surface of the cuvette must be protected from damage. Do not touch the surface with fingers or rough things. After assay, the cuvette must be washed clean. If you are having trouble removing residues from cuvettes, try cleaning with ethanol or methanol. They often works well to remove contaminating proteins. A thorough wash with distilled water is always a must, do not remain any measuring solution in the cuvette.
4. Light detecting system
This system can turn the light energy into the electric energy and the result can be measured on the galvanometer and shown on its display.
Ⅲ
1.Turn the instrument on and the power light comes on. Allow the instrument to warm up for approximately 20 min before use.
2. Turn the wavelength selector knob to the desired wavelength.
3.Use [MODE] button setting test manner: Transmittance(T), Absorbance(A), Concentration(C) ect. We usually select Absorbance(A) MODE.
4.Insert the the cuvettes containing the reference (blank) solution and the sample solution in the holder in order. Then close the lid of the holder.
5. Put the cuvette containing the reference solution into the light path, then press “100%” button. Now the display shows “BLA” and we should wait a minute until it shows “0.000”A.
6.When the display shows “0.000”A , Set the cuvette to be read in the light path by pulling the pulling rod, read and write down the absorbance shown by the display.
7. After finishing the reading of all the samples, take out the cuvettes and lid the sample compartment, switch off the power supply. The cuvettes must be rinsed first with tap water thoroughly, and then with distilled water and dried before use.
Spectrophotometer is a sophisticated apparatus, so care must be taken when using it. It must be protected from shock, dump and corrosion.
The condition of the glass surfaces of the cuvettes is closely related to the absorbance.

Make sure cuvettes are clean of all residues and in order to keep them clean and transparent, the cuvettes must not be touched by fingers and hard things(e.g., tube brush). Avoid filling the cuvette too much to prevent corrosion of the cuvette holder or the sample compartment.
Always be sure that the solutions are well mixed before measurements are made.
In a biochemical laboratory, the protein assay is a commonly performed assay yet is often weakly understood. The value of the protein assay is vital since it is usually used to decide another value of biochemical determination. Errors in protein determination tend to amplify overall errors in laboratory calculation. There are several kinds of methods to determinate protein concentration, and each method has different advantages and limitations. Before starting a particular analysis, it is a good idea to be aware what other researchers in you field are utilizing to determine protein concentration. Then we can select better method by means of theirs. All of the described methods rely on the use of a spectrophotometer. So before beginning these assays you should make yourself familiar with the operation and use of a spectrophotometer.
The objective of this experiment is to apply the principle and technique of spectrophotometry to the determination of proteins.
1.Become familiar with spectrophotometry
2.Master the method of determination of protein
3.Learn to make standard curve
Coomassie brilliant Blue G-250 has 2 forms of color: brown and blue. This dye can specifically bind to proteins at arginine, tryptophan, tyrosine, histidine and phenylalanine residues. After it binds with protein, the brown color will turn into blue, which has an absorbance maximum at 595 nm (blue). The concentration of protein in solution is proportional to the depth of blue color, but the assay does perform linearly only over low concentration stretches. In this way, we can calculate the protein with the help of 2000 model spectrophotometer.
1. Pick up 3 tubes, add in reagents as the table below:
Reagents(ml) Control tube Standard tube Assay tube

0.9% NaCl standard protein (50 μ g/ml)
Sample protein
Coomassie blue reagent
0.1
–
–
5.0
–
0.1
–
5.0
–
–
0.1
5.0
2. Mix each tube sufficiently, then at room temperature for 5 minutes, then set the solution to glass cuvette, with the help of 2000 model spectrophotometer (λ=595nm), adjust “0”with solution in control tube, read the A value of solution of assay and standard tube respectively and record.
3. Calculate : sample protein concentration( μ g/ml)=A of assay tube solution/A of standard tube solution ×
50( μ g/ml)
1.
The dye binds to quartz cuvettes so it is usually better to use glass cuvettes. If you have trouble removing residues from cuvettes, try cleaning with ethanol or methanol. They often works well to remove contaminating proteins.
2.
The assay does perform linearly only over low concentration stretches.
Determination of protein concentration by ultraviolet absorption (280 nm) depends on the presence of aromatic amino acids in proteins. Tyrosine and tryptophan absorb at approximately
280 nm. Protein also has the character of ultraviolet absorption. The highest value is in the region of 280nm. The concentration of protein in solution is proportional to the A value. In this way, we can calculate the protein with the help of UV model spectrophotometer. The real advantages of this method of determining protein concentration are that the sample is not destroyed and it is also very rapid. Although different proteins will have different amino acid compositions and thus different molar absorptivities, this method can be very accurate when comparing different solutions of the same protein.
Ⅰ . Plot the standard curve
（ 1 ） Standard protein solution: its concentration is 1mg/ml.
（ 2 ） Pick up 6 tubes, add in reagents as the table below:
Reagents(ml) 1 2 3 4 5 6

Standard protein solution
0.9% NaCl
0.5
3.5
1.0
3.0
1.5
2.5
2.0
2.0
2.5
1.5
0
4.0
（ 3 ） Mix each tube sufficiently, then set the solution to quartz cuvette, with the help of UV spectrophotometer (λ=280nm), adjust “0”with solution in the 6 th tube, read the A value of solution of the front 5 tubes respectively and record.
（ 4 ） Place the protein concentration on the y-axis and the A on the x-axis, plot the standard curve.
Ⅱ. Determine the sample protein solution
（ 1 ） Pick up 2 tubes, add in reagents as the table below:
Reagents(ml)
Sample protein solution
Control tube
0
Assay tube
1.0
0.9% NaCl 4.0 3.0
（ 2 ） Operate as before, adjust “0”with solution in control tube, read the A value of solution of assay tube.
（ 3 ） Refer the standard curve, protein concentration can be found according to A value of the solution in assay tube.
The biuret reaction is a method that can be used to determine the amount of protein in a solution. The biuret reagent (copper sulfate in a strong base) reacts with peptide bonds and forms red-purple product when it does so. The spectrophotometer can then be used to measure the intensity of the color produced. The more protein present, the deeper the color. The concentration of protein in solution is proportional to the depth of color. In this way, we can calculate the protein amount with the help of model 2000 spectrophotometer.
Plot the standard curve
1. Standard protein solution: its concentration is 60mg/ml.
2. Use your brain actively, design an experiment (biuret method to determine protein) to produce a standard curve according to your results. Pick up 6 tubes, add in a series of serial standard protein dilutions and biuret reagent 4ml per tube.
3. Mix each tube thoroughly but gently and allow them to set at 37 ℃ water for 15 minutes.
Then set the solution to glass cuvette respectively, with the help of model 2000 spectrophotometer
(λ=520nm), adjust “0”with blank solution in the tube, read the absorption (A) value of solution of other tubes respectively and record.
4. Plot a standard curve with the amount of protein in the test as the abscissa, and the absorption as the ordinate.

1.The red-purple compound has an absorbance maximum at 520 nm.
2.The concentration of protein in solution is proportional to the depth of red-purple color, but the assay does perform linearly only during 1~20mg/ml concentration stretches.
3.The sample should be diluted before they are tested. Also different samples may need to be diluted very differently. One will hopefully fall within the range of the standard curve.
Here is an example of how to do a series of serial dilutions :
1 ml extract + 4 ml water = 1/5 dilution
1 ml 1/5 dilution + 4 ml water = 1/25 dilution
1 ml 1/25 dilution + 4 ml water = 1/125 dilution
Tips for Success
1. Write down a clear plan before you start working.
2. Keep very careful notes so you can recheck what you did if you get unexpected results.
3. Label all tubes and samples so they don't get mixed up.
4. Work slowly and carefully. Accurate, reproducible results in a biochemical test requires care on your part. If you are sloppy or careless your results will suffer.
(Cui Fu-ai)

Chromatography is used to separate or to analyze complex mixtures. It is based on differential migration. Since the various components in a mixture have different relative chemical properties, such as molecular size, shape, polarity, binding specificity, charges, affinity, et al, when they pass through a stationary phase, the different components have different migration speed and therefore they will be separated from each other.
In all chromatography, there is a mobile phase and a stationary phase. The stationary phase is the phase that doesn't move and the mobile phase is the phase that does move. (Although the stationary phase may appear to be a solid phase, in many cases it is actually a liquid stationary phase.) The mobile phase is normally allowed to pass over the stationary phase. As the mobile phase travel through the stationary phase, it will pick up the compounds to be separated with it. At different points the different components of the compounds are going to be absorbed and are going to stop moving with the mobile phase due to their different chemical properties. Those molecules which are absorbed only weakly with the stationary phase will spend much shorted time in this phase and therefore will be eluted first, whereas those which interact most strongly with the stationary phase will spend more time in this phase and will be eluted later. Sometimes it is necessary to change the composition of the buffer during the elution process to remove the most strongly absorbed molecules.
There are several types of chromatography used in biochemical analysis, including paper chromatography, thin-layer chromatography, column chromatography; gas chromatography (GC), liquid chromatography (LC); ion exchange chromatography, gel chromatography, affinity chromatography et al.

Paper chromatography: In paper chromatography the mobile phase is the solvent. The stationary phase is the strip or piece of paper that is placed in the solvent. This kind of chromatography use capillary action to move the solvent through the stationary phase. A small spot of the solution containing the sample is applied to a strip of chromatography paper about one centimetre from the base. This sample is adsorbed onto the paper. This means that the sample will contact the paper and may form interactions with it. Any substance that will react with (and thus bond to) the paper cannot be measured using this technique. The paper is then dipped into a suitable solvent (such as ethanol or water) and placed in a sealed container. As the solvent rises through the paper it meets the sample mixture which starts to travel up the paper with the solvent.
Different compounds in the sample mixture travel different distances according to how strongly they interact with the paper. Paper chromatography takes some time and the experiment is usually left to complete for some hours.
Column chromatography: The stationary phase is a powdered adsorbent which is placed in a vertical glass column. The mixture to be analysed is loaded on top of this column. The mobile phase is a solvent poured on top of the loaded column. The solvent flows down the column, causing the components of the mixture to distribute between the powdered adsorbent and the solvent, thus (hopefully) separating the components of the mixture so that as the solvent flows out of the bottom of the column, some components elute with early collections and other components elute with late fractions.
Thin Layer chromatography: In thin-layer chromatography the mobile phase is the solvent.
The stationary phase is a powdered absorbent which is fixed to a aluminum, glass, or plastic plate.
Like paper chromatography, thin Layer chromatography also use capillary action to move the solvent through the stationary phase. The mixture to be analyzed is loaded near the bottom of the plate. The plate is placed in a reservoir of solvent so that only the bottom of the plate is submerged.
This solvent is the mobile phase; it moves up the plate causing the components of the mixture to distribute between the adsorbent on the plate and the moving solvent, thus separating the components of the mixture so that the components are separated into separate "spots" appearing from the bottom to the top of the plate.
Gel filtration chromatography: It is also called molecular exclusion or gel permeation chromatography. Gel filtration chromatography is a separation based on size. The stationary phase

consists of porous beads with a well-defined range of pore sizes. The stationary phase for gel filtration has a fractionation range, meaning that molecules within that molecular weight range can be separated. The liquid phase consists of an aqueous buffer solution. A mixture of macromolecules is added to the top of a column filled with Sephadex and buffer. As the macromolecules move through the buffer, smaller molecules become trapped in the pores, and take longer to elute. Larger molecules don’t fit into the pores as well and elute more quickly. Very large molecules don’t fit into the pores at all and move between Sephadex beads, thus eluting the quickest. This principle can be utilized for separating molecules according to molecular weight, although there are limits for molecular weight ranges that are suitable for a particular formulation.
Gel filtration can also be used to remove salts (desalting) and other low molecular weight contaminants from protein solutions. Proteins that are small enough can fit inside all the pores in the beads and are said to be included. These small proteins have access to the mobile phase inside the beads as well as the mobile phase between beads and elute last in a gel filtration separation.
Proteins that are too large to fit inside any of the pores are said to be excluded. They have access only to the mobile phase between the beads and, therefore, elute first. Proteins of intermediate size are partially included - meaning they can fit inside some but not all of the pores in the beads.
These proteins will then elute between the large ("excluded") and small ("totally included") proteins.
The space between gel beads is called the void volume, Vo. The space within the beads is called the internal (pore) volume, Vi. The total volume of mobile phase in the column will be Vt, or the sum of Vo and Vi. Large molecules that cannot pass through the gel pores will be excluded from the gel beads and pass through the column in the void volume, Vo. Molecules much smaller than the pore size will equilibrate with the entire liquid volume and elute at a volume equal to Vt.
Molecules with intermediate size will pass through some of the pores but not all pores and elute at some volume, Ve (elution volume) in between the two extremes, Vo and Vt, depending on how large they are and what fraction of pores in the beads they can pass through. A plot of retention volume vs log MW will generally give a straight line, although deviations from linearity are not uncommon, due to differences in protein shapes (not all are spherical).
Ion exchange chromatography: Ion exchange chromatography separates molecules based on their electrical charges which depend on their amino acid composition and the pH of the medium.

It relies on charge-charge interactions between the proteins in your sample and the charges immobilized on the resin of your choice. In this type of chromatography, the resin (the stationary solid phase) is used to covalently attach anions or cations onto it. Solute ions of the opposite charge in the mobile liquid phase are attracted to the resin by electrostatic forces.
In ion exchange chromatography, charged substances are separated via column materials that carry an opposite charge. The ionic groups of exchanger columns are covalently bound to the gel matrix and are compensated by small concentrations of counter ions, which are present in the buffer. When a sample is added to the column, an exchange with the weakly bound counter ions takes place. Since the bound biomolecules are subsequently displaced with the aid of an increasing salt gradient protein varying in charge can be separated. The desorption of the proteins from the column begins only with steadily increasing ionic strength of the elution solution or pH changes that can be modified as to give your protein or the matrix a charge at which they will not interact and your molecule of interest elutes from the resin. The substances that have a higher charge density, are bound correspondingly stronger to the column while the others elute rapidly.
There are two types of exchange: cation exchange in which the stationary phase carries a negatively charged groups which bind positive site on the protein, and anion exchange in which the stationary phase carries positively charged groups which bind negative site on the protein. In cation-exchange chromatography, the rate of mobility of proteins loaded onto the resin is proportional to the degree of negative charge that they bear. That is to say, proteins with a more negative net charge will move faster and elute easier. On the contrary, in anion exchange chromatography, proteins with a more positive net charge will move faster and elute easier.
Therefore, if we want to get positive net charge first, we may use anion exchange chromatography.
Affinity chromatography ： In Affinity chromatography, proteins are separated according to their ability to bind to a specific ligand that is connected to the beads of the resin. After the proteins that do not bind the ligand are washed through the column, the bound protein of interest is eluted from the column by washing with increasing concentrations of free ligand. Because the interactions in this type of chromatography are so specific it affords the greatest degree of purification in a single step. The drawback is the need to make individual column for each molecules to be purified.

In this experiment we will be using two kind of liquid chromatography, also known as column chromatography because it uses large open columns. They are gel filtration chromatography and ion exchange chromatography.
There are five proteins in serum. They are albumin, α
1
-globulin, α
2
-globulin, β -globulin and γ -globulin. In this experiment, our aim is to get purified γ -globulin and remove the other four proteins from serum. We use three methods: salting out, gel filtration chromatography and ion exchange chromatography.
Ⅰ .Crude extract — salting out: The aim of this step is to remove albumin from the serum.
This steps involves purifying proteins based on their solubilities in varying concentrations of ammonium sulfate. The solubility of proteins depends on the relative balance between protein-solvent interaction, which keep it in solution, and protein-protein interactions, which cause it to aggregate and precipitate. Low-ionic strength (low ammonium sulfate concentration) can enhance protein-solvent interaction, protein remain in the solution. That is to say, the solubility of a protein at low ionic strength generally increases with the salt concentration (salting in). High ionic strength (high ammonium sulfate concentration) can enhance protein-protein, interaction, protein aggregate and precipitate. That is, the solubility of a protein at high ionic strength decreases (salting out). We can make protein precipitate by increase salt ionic strength. Since different proteins have different sizes and charges, they precipitate at different salt concentration.
We can adjust the salt concentration in a solution to just over the precipitation point of the protein of interest. In this case, the protein of interest will be precipitated while many other will remain in solution. In this experiment, the albumin molecules are small and have more charges. Only in saturated ammonium sulfate solution can it precipitate. While globulin molecules are bigger than

albumin and have much fewer charges, they can form precipitation in semi- saturated ammonium sulfate. So we can adjust the concentration of ammonium sulfate to be semi-saturated. Then globulin will precipitate, while albumin still remains in solution. By this way, we can separate albumin and globulin.
Ⅱ .Desalt
— Gel filtration chromatography: After we finished the step above, the globulin solution would contain a lot of ammonium sulfate. The aim of this step is to desalt. We use gel filtration chromatography. Gel filtration chromatography separate molecules according to their different size. We all know that the molecules of ammonium sulfate are much smaller than proteins. Therefore, ammonium sulfate will take longer time to elute from the column than proteins. That is to say, proteins can be eluted faster than the ammonium sulfate. We can get globulin first. By this way, the salt of ammonium sulfate will be removed from the globulin solution.
Ⅲ .Purify of γ -globulin — Ion exchange chromatography: This step is to remove the other three globulin and obtain γ -globulin. In this experiment, we use anion ion exchange chromatography. The pH of elution buffer is 6.5.The stationary phase is DEAE cellulose in which carrier positively charged groups. The pI of α
1
, α
2
, β -globulin is less than 6.0.They will carry negative charge in the pH6.5 elution buffer. While the pI of γ -globulin is 7.3. It will carry positive charge in pH6.5 buffer. In anion exchange chromatography, protein with positive net charge will move faster and elute easier. So we can get γ -globulin first and separate γ -globulin and the other three globulins.
1. Salting out
Take 1.0ml serum into a test tube. Then add 1ml saturated (NH
4
)
2
SO
4
solution into the tube slowly and shake the tube from time to time. Lay the tube aside at room temperature for about 10 minutes. Then centrifuge 3000rpm, 10mins. Discard the supernate that contain albumin and remain the sediment that contain four globulins. Then add 0.5ml water to the sediment and make the sediment dissolve. [Note: When you use centrifuge, make sure balance the load in the rotor-every tube must have a balance tube in the opposite slot with the same volume of fluids.
2. Desalt- Gel filtration chromatography
① Equilibrate the column with elution buffer by passing about the buffer through the

column bed after it has completely settled. [Note: the Sephadex gel has already been soaking in the elution buffer]. Avoid stirring up the top of the column bed when adding buffer or samples, as this will give poor resolution of samples. Be especially careful not to allow the top of the gel to dry. Always keep buffer solution above the top of the gel in the column.
② Allow the buffer solution to drain down until there is only about 1 or 2 mm of buffer above the gel before turning off the stopcock. Then carefully add the protein sample mixture to the top of the column using a pasteur pipet. Again, do not stir up the top of the gel. Open the stopcock to allow the protein solution to enter the gel bed but not to make the top of the gel dry.
③ Carefully add elution buffer to the top of the column. Replenish the buffer in the column throughout the experiment.
④ In the process of elution, you should examine the outflow of protein from time to time.
Take a drop of effluence in a glass plate and add a drop of sulfonic acid. When the effluence in the glass plate become turbid, begin collecting fractions in a tube. In the process of collection, you also need examine the effluence from time to time. When the effluence is no longer become turbid, stop the collection .At no time should the level of the buffer drop below the level of the packed
Sephadex.
3. Ion exchange chromatography:
The process is all the same to the above.
4. Condense
Add Sephadex G-25 to the collection fractions until there is only about 1 or 2 mm liquor level.
γ
—
To identify the protein that we have extracted from serum.
Molecules with a net charge (like protein, nucleic acid) will move in an electric field - this is called electrophoresis. Electrophoresis is a common method of separating macromolecules (such as proteins, DNA and RNA). The separation relies on the different rates of movement of molecules in electric fields. The velocity of migration of charged molecules will depend upon the strength of the electric field, the net charge on the molecule and the size ， shape of the molecule.

Small molecules move faster than large molecules. Globular molecules move faster than elongated fibrous molecules. The greater charges the molecules have, they will move faster. Since different molecules have different molecules size, shape and charges, they will migrate at the different rate in an electric fields. So they can be separated from each other.
In this experiment, we will use cellulose acetate membrane electrophoresis to identify the protein that we have extracted from serum. We will make serum electrophoresis and the protein we have extracted from serum electrophoresis at the same time. We all know that there are five kinds of proteins in serum. The pI of all these five proteins are less than 8.6. When we use pH8.6 barbital as the electrophoresis buffer, all the protein will carry net negative charges. They will migrate to the positive pole of the electric field. Since the size of albumin molecule is the smallest and the net charges on the albumin molecule are the most in all five serum proteins, the migration speed of the albumin molecule is the fastest, and then α 1-globulin, α 2-globulin, β -globulin, the slowest is γ -globulin. If the protein that we have extracted from serum has the same migration rate with the γ -globulin in serum, it is clear that we have got the protein we want.
1. Add the sample to the membrane:
①
Take out the membrane from the barbital buffer.(Each group take two membrane, one for serum, and another for the protein that we have extracted).
② Absorb superfluous water in the surface of membrane with filter paper. Be especially careful not to allow the membrane to dry.
③ Draw a clear line that keep 1.5cm distance from one end of the membrane with a pencil. ④
Take out a filter paper slice with a nippers and put the filter paper slice into the sample (serum or the extract solution). Put the filter paper slice that have absorbed the sample to the sample line in a vertical way.
⑤ Put the membrane to the negative pole of the electrophoresis slot and make sure that the membrane surface containing sample is downward.
2. Turn on the electric power: Adjust the voltage to 90-110v. Power on about 45-60min. Then turn off the power and take off the membrane from the electrophoresis slot.
3. Stain the membrane: Stain the membrane in the staining agent for about 2-5min.

4. Bleach the membrane: Bleach the membrane in the bleaching liquor until the background become white and the protein bands are very clear.
(Liu Zhi-fang)

A great deal of knowledge can be obtained by observing the behavior of charged molecuels traversing a uniform electric field. To obtain a uniform electric field with a constant magnitude and direction over a specified volume of space, two flat metal plates are set up parallel to each other as shown in Fig. 1. When the terminals of a power source with voltage V are connected to these plates, as indicated in the diagram, a uniform electric field E is produced between the plates.
Out side of the plates and near the ends, the field is not uniform.
Electrophoresis is the migration of charged molecules like proteins in an electrical field.
The separation of proteins in an electric field is based on the size, shape, and charge. The charges are contributed by the side chains of the amino acids of which the proteins are composed. The charge of the protein depends on the IpH of the protein and the pH of the surrounding buffer.
Most electrophoretic methods use a supporting media, such as starch, paper, polyacrylamide, or agarose. The term zone electrophoresis refers to electrophoresis which is carried out in a supporting medium, whereas moving-boundary electrophoresis is carried out entirely in a liquid phase. When proteins are visualized on gels and the migration distances are compared to standards the isoelectric pH (isoelectric focusing) and molecular weights (SDS electrophoresis) of various proteins can be measured. The isoelectric pH and molecular weights are uselful in identifying and purifying proteins.
When you apply a force (F=MA) to a mass, the mass will accelerate(A). Acceleration is defined as the rate of change in velocity as shown in Equation 1.

Equation 1 :
A = (v-vo)/T
A = acceleration v = final velocity v o
= initial velocity T = time
The acceleration of the spherical molecule will be zero when the drag force equals the accelerating force (Equation 2.). The molecule will move with a constant velocity.
Equation 2:
Fa = Fd
The driving force of a molecule in an electrical field is equal to electrical field strength (E) times (multiply) the charge of the molecule (Q). The equation of electrical field strength is shown in Equation 3. The electrical field strength (E) has units of volts/cm and the charge (Q) has units of coulomb. The rate of migration of an ion, therefore, can be altered by changing the distance between electrodes as well as by changing the electrical potential (V). In the absence of a resisting force, ions would accelerate in an electrical field. They assume a constant velocity because of the resistance.
Equation 3 :
Fa = E Q
E = (V/d)
V = volts d = cm Q = coulomb
The resisting force is a function of size, shape, and viscosity in accordance with the Stokes equation. The drag force is the retarding force as the protein accelerates through the gel. It is proportional to the frictional coefficient and the velocity of the protein molecule (Equation
5).When the resisting force exceeds the driving force, no ion migration takes place. When the driving force exceeds the resisting force, there is an accelerated migration. But when a constant velocity is achieved, the accelerating force (Fa) is equal to the resisting force (Fd).
Equation 4 :
Fd = 6
π r
η v
η = viscosity v=velocity
When F a
= F d
, then
Equation 5:
EQ = 6
π r
η v

Equation 6 :
µ = V/E = Q/6 π r
η
Since µ is the mobility and is the rate of migration of a charged molecule in a field of unit strength (if E=1 volt/cm), then the velocity (v) becomes mobility (µ ). The electrophoretic mobility is directly proportional to net charge (Q) and inversely proportional to the size of the molecule(r) and to the solution viscosity ( η ) as shown in Equation 6.
The mobility of a protein in an electrical field is defined as the velocity (V) of the protein molecule per electrical field strength (E). In a fix field strength, the mobility (µ ) of a protein is proportional to the charge(Q) and inversely proportional to the size of the molecule and the viscosity of the buffer. The type of buffer used and the pH of the buffer are important, since the net charge of the protein is determined by the pH of the buffer. The charge of a protein depends on the ionization of the acidic and basic groups on the protein, which is a function of the pH of the medium.
The binding of buffer ions other than H+ by protein molecules is important. When quoting a value for electrophoretic mobility, it is imperative to state the conditions of the buffer, such as pH, ionic strength, and buffer composition. It has become customary to carry out electrophoresis of serum proteins at a pH of 8.6. At this pH, 85% of all proteins are negatively charged and will migrate to the anode.
Gel electrophoresis is a technique used for the separation of nucleic acids and proteins.
Separation of large (macro) molecules depends upon two forces: charge and mass. When a biological sample, such as proteins or DNA, is mixed in a buffer solution and applied to a gel, these two forces act together. The electrical current from one electrode repels the molecules while the other electrode simultaneously attracts the molecules. The frictional force of the gel material acts as a "molecular sieve," separating the molecules by size. During electrophoresis, macromolecules are forced to move through the pores when the electrical current is applied. Their rate of migration through the electric field depends on the strength of the field, size and shape of the molecules, relative hydrophobicity of the samples, and on the ionic strength and temperature

of the buffer in which the molecules are moving. After staining, the separated macromolecules in each lane can be seen in a series of bands spread from one end of the gel to the other.
There are two basic types of materials used to make gels: agarose and polyacrylamide.
Agarose is a natural colloid extracted from sea weed. It is very fragile and easily destroyed by handling. Agarose gels have very large "pore" size and are used primarily to separate very large molecules wiht a molecular mass greater than 200 kdal. Agarose gels can be processed faster than polyacrylamide gels, but their resolution is inferior. That is, the bands formed in the agarose gels are fuzzy and spread far apart. This is a result of pore size and it cannot be controlled. Agarose is a linear polysaccharide (average molecular mas about 12,000) made up of the basic repeat unit agarobiose, which comprises alternating units of galactose and 3,6-anhydrogalactose. Agarose is usually used at concentrations between 1% and 3%. Agarose gels are formed by suspending dry agarose in aqueous buffer, then boiling the mixture until a clear solution forms. Then poured and allowed to cool to room temperature to form a rigid gel.
The polyacrylamide gel electrophoresis (PAGE) technique was introduced by Raymond and
Weintraub (1959). Polyacrylamide is the same material that is used for skin electrodes and in soft contact lenses. Polyacrylamide gel may be prepared so as to provide a wide variety of electrophoretic conditions. The pore size fo the gel may be varied to produce different molecular seiving effects for separating proteins of different sizes. In this way, the percentage of polyacrylmide can be controlled in a given gel. By controlling the percentage (from 3% to 30%), precise pore sizes can be obtained, usually from 5 to 2,000 kdal. This is the ideal range for gene sequencing, protein, polypeptide, and enzyme analysis. Polyacrylamide gels can be cast in a single percentage or with varying gradients. Gradient gels provide continuous decrease in pore size from the top to the bottom of the gel, resulting in thin bands. Because of this banding effect, detailed genetic and molecular analysis can be performed on gradient polyacrylamide gels. Polyacrylamide gels offer greater flexibility and more sharply defined banding than agarose gels. Different samples are loaded in wells or depressions at the top of the polyacrylamide gel. The proteins move into the gel when an electric field is applied. The gel minimizes convection currents caused by small temperature gradients, and it minimizes protein movements other than those induced by the electric field. Proteins can be visualized after electrophoresis by treating the gel with a stain such

as Coomassie blue, which binds to the proteins but not to the gel itself. Each band on the gel represents a different protein (or a protein subunit); smaller proteins are found near the bottom of the gel.
Isoelectric electrophoretic focusing (IEF) is an excellent method for the analysis and preparative isolation of proteins. It is also ideal for determining purity. It has excellent resolution down to .02 pH units. IEF gives more separation than any other method. When an protein that appears homogeneous by other methods produces several bands by IEF, this could be due to better separation abilities by IEF meaning the protein was not pure. It could also be due to the presence of isomers, differences in primary structure, kinds and numbers of prosthetic groups, or the denaturation of the protein. Elution of proteins from a gel is a method of obtaining separated or purified proteins for other assays such as sequencing and tryptic digestion.
[principles] The structure, molecular size and charge of all kinds of lipoprotein are different, so they can be separated in a uniform electric field. According to the electrophoresis rate, plasma lipoprotein can be divided to four parts: α lipoprotein ， preβ lipoprotein, β lipoprotein and

CM. CM is chylomicron and preβ lipoprotein belong to VLDL, β lipoprotein belongs to LDL, and α lipoprotein belongs to HDL.
[methods]
1. pre-dyeing of serum: obtain 1ml serum from ear vein of rabbit, then add 0.5 ml Sudan Black.
Put the mixture in 37 ℃ water for 30min, then centrifuge at 2000rpm for 5min. The supermatant is used for the next step.
2. prepare agarose gel: Put a clean slide on the desk, then add 3.5ml melted 0.5% agarose gel
(buffer: pH8.6 barbiturate) on it, waiting for its solidify.
3. add sample: draw 0.02ml pre-dyed serum ,then add it on the two piece filter paper (L:1.2cm, w: 0.2cm). Insert the paper to the coagulate gel and the distance is 2cm.
4. electrophoresis: voltage -100V, time-1hr. Then observe the result.
[Note]
1.
the melted gel must be once added.
2.
gel must be put for at least 30min till it is coagulate.
[principle] Insulin is normally secreted by the beta cells (a type of islet cells) of the pancreas. It can regulate the level of blood glucose. When glucose is boiled with high concentration acid in
100 ℃ water, the product can react with O-Toluidine and the color of react solution becomes green. The colors suppose the level of blood glucose.
[methods]
1. blood getting: (A) Get 2 ml blood from ear vein .of rabbit. (B) after injection of 1 ml insulin for
40min, get 2 ml blood as before.
2. centrifuge the two sorts of blood samples above at 4,000 rpm for 10 min, and get the serum.
3. take 4 tubes and operate as below.
Reagent(ml)
Serum
Standard (1mg/ml)
Tube(A)
0.1
Tube(B)
0.1
Standard
0.1 control

ddH2O 0.1
O-Toluidine 4.0 4.0 4.0
4. boil the mixture in 100 ℃ water for 10 min, then take in cold water at once.
4.0
5. make the OD of control zero and measure the OD of others.
6. calculation:
(OD of samples/OD of standard) × 1mg/ml × 1000 ÷ 180=blood glucose mmol/L
7. To compare the blood glucose levels in the two sorts of blood samples with and without insulin injection.
[Note]
1.Do not get too much blood at the first time.
2.insulin is injected from subcutaneous.
[principle] In the process of glycolysis, the further reaction of two triose phosphate need key enzyme glyceraldehydes-3-phosphate dehydrogenase. The –SH is the essential group of this enzyme, so if the inhibitor of –SH added, glycolysis will stop at the stage of triose phosphate, and triose phosphate will be accumulated. Triose phosphate can react with 2,4-Dinitrophenylhydrazine ， and the products appear mauve.
[methods]
1. mix 1g yeast and 50 ml 5% glucose liquor together.
2. take three tubes, and operate as below:
Reagent (ml) pH7.4 phosphate buffer
CH
2
ICOOH
Hydrazine sulfate
10%CCl
2
COOH yeast liquor
H
2
O
CH2ICOOH
Tube A
2.0
0.5
0.5
1.0
Tube B
2.0
0.5
0.5
—
4.0 4.0
— 1.0
Keep the mixtures in 37 ℃ water for 1hr.
— —
Tube C
2.0
—
—
—
4.0
2.0
0.5

Hydrazine sulfate
10%CCl
2
COOH
—
—
2.0
—
1.0
H
2
O 1.0
3. After 10min, centrifuge the mixtures at 3000rmp for 10min,
4. After get the supernatant above, take 3 new tubes and operate as below:
0.5
1.0
—
Reagent (ml) supernatant
1mol/LNaOH
Tube 1
0.5 (Tube A)
Tube 2
0.5 (Tube B)
0.5 0.5
Keep the mixtures in room temprature for 10min.
Tube 3
0.5 (Tube C)
0.5
2,4-Dinitrophenylhydrazine
1mol/LNaOH
0.5
3.5
0.5
3.5
5. Analyze the results.
[Note]
1. The experiment needs many reagents, so it is necessary to see the tags clearly.
2. The reagents need correctly quantity to undertake the results.
(Zeng ji-ping)
0.5
3.5

In the biotechnology industry, we are often faced with the problem of separating various biomolecules (from large organisms and cells to proteins) . Centrifugation is often a preferred method for conducting this job, due to the simplicity and reliability of the process. It separates particles from a solution according to their sizes, shapes, relative densities, viscosity of the medium and rotor speed. While there are numerous types of centrifuges with varied configurations, all operate on the same basic principle of applying a physical force to an object and causing it to move in a medium. The physical force is called centrifugal force .It is to say: a particle in a centrifugal field will experience a centrifugal force defined by: Fc = m ω 2 r .
Where Fc = the centrifugal force, m = mass of the particle,
ω
= angular velocity r =distance from the axis.
We can see that the centrifugal force generated is proportional to the rotation rate of the rotor
(in rpm)and the distance between the rotor center and the centrifuge tube and the mass of a particle. Usually we also use relative centrifugal force (RCF). RCF is defined as the ratio of the centrifugal force to the force of gravity: RCF = Fc/Fg = ω 2 r/980. ω (expressed in radians/sec) is converted to revolutions per minute (rpm) by substituting ω = π (rpm)/30, resulting in: RCF =
1.119 x 10-5(rpm)2r here r is expressed in cm. RCF units are expressed as "x g".
The centrifuge is an essential instrument in cell and molecular biology research. It is primarily used to separate biological components based upon differential sedimentation properties. Many types of centrifuges are available for various applications. All centrifuges basically consist of a motor which spins a rotor containing the experimental sample. The rotor spins when the motor is turned on, which generates a centrifugal force that causes the targeted particles to sediment out in the bottom of the container. The differences between centrifuges are in the speeds at which the samples are centrifuged and the volumes of samples.
①

The two most common type of centrifugation are analystical and preparative; the distinction between the two is based on the purposes of centrifugation .Analytical centrifugation involves measuring the physical properties of the sedimenting particals such as sedimentation coefficient or molecular weight. Optical methods are used in analytical centrifugation. Molecules are observed by optical system during centrifugation, to allow observation of macromolecules in solution as they move in gravitational field.
Preparative centrifugation is used most often in the biological sciences to collect material or to separate particles based upon their sedimentation properties. Preparative centrifugation takes advantage of the fact that more massive particles will sediment faster than less massive particles.
For example, organelles and other subcellular components can be isolated by differential centrifugation. Differential centrifugation is carried out by centrifuging a sample at low speed and separating the supernatant and pellet. The supernatant is then recentrifuged at higher speed and the supernatant and pellet separated again. Sedimentation of a particular particle is dependent upon the RCF and how long the centrifugal force is applied. Samples must be centrifuged long enough for the particles at the top of the tube to reach the bottom. The size and shape of the centrifuge tube will also affect the centrifugation time. In addition, there is a difference in the RCF between the top and bottom of the tube. These factors need to be taken into consideration when separating a component. The term g.min refers to the product of the minutes a sample is centrifuged and the g-force. In general, assuming that the sizes and dimensions of the centrifuge tubes are similar, the product of the g-force and the time of centrifugation can be used to determine the conditions needed to completely sediment a particle. For example, if it takes 60 min to completely sediment a particle at 1000xg, then it should only take 10 min to sediment the same particle at 6000xg.
②
Fast sedimenting particles will be contaminated with slow sedimenting particles. The reason for this is that by the time large particles near the top of the tube are pelleted, some of the small particles near the bottom of the tube have also pelleted. In addition, mechanical vibrations, thermal gradients and convection currents can also affect the sedimentation properties.
Centrifugation through a dense medium, or density gradient centrifugation, can partially alleviate

these problems. In addition, density gradient centrifugation will allow for better separation of particles with similar properties. Several different media are commonly used in density gradient centrifugation. Preforming the gradients before centrifugation decreases the amount of time and centrifugal force needed to reach equilibrium. The two types of density gradient centrifugation are rate zonal and isopycnic (or equilibrium).
In rate zonal centrifugation the density of the particles being separated are greater than the density of the solvent. Separation is based primarily upon size (i.e., larger particles will sediment faster). It is important to determine the optimal length of centrifugation for separating the particle of interest. If the centrifuge is not turned off soon enough all of particles will pellet. In the rate zonal density gradients it is necessary to use preformed gradients. Gradient makers are used to produce continuous gradients, or stepwise gradients can be prepared by layering successive solutions of lesser density. Samples are layered onto the gradients and subjected to centrifugation.
In isopycnic centrifugation the solvent density encompasses density of particles. The separation is based upon particle density. Centrifugation is carried out until equilibrium is reached
(i.e., all particles have banded at densities corresponding to their own). In isopycnic density gradients it is possible to mix the sample with the density gradient medium and carry out the centrifugation until the density gradient forms. The various components in the sample will then be found at a position which corresponds to their density. This is especially useful in the case of
Percoll which rapidly forms density gradients when subjected to centrifugation. Another common example in which self-forming gradients are commonly used is the separation of nucleic acids on
CsCl gradients. In isopycnic gradients the sample can also be underlaid at the bottom of the tube and the various particles will 'float' to their correct densities during centrifugation. Following centrifugation the gradient is divided into fractions corresponding to different densities and analyzed.
③
Another system of classification is the rate or speed at which the centrifuge is turning.
Ultracentrifugation is carried out at speed faster than 20,000 rpm. Super speed ultracentrifugation is at speeds between 10,000 and 20,000. Low speed centrifugation is at speed below 10,000 rpm.
① Make sure the correct rotor is being used and that it is installed properly on the spindle.

② Balance the load in the rotor-every tube must have a balance tube in the opposite slot with the same volume of fluids.
③ Make sure you are using the appropriate centrifuge tube for the job-they can rupture at too high a speed. smoothly.
Ⅰ
Nucleic acid is a kind of important biomacromolecules. They include DNA and RNA. In vivo, DNA and RNA usually bind with different protein and form nucleoprotein. That is to say, they exist in the form of DNP and RNP. In this experiment, our aim is to abstract RNA from the animal liver cells. First of all, we should breakdown the cells and prepare liver homogenate. We can use homogeniter to make the cell membrane rupture and release nucleic acids. Second, according to the difference of solubility in salt solution of RNP and DNP, we separate DNP and
RNP. RNP can easily dissolve in 0.14mol/L NaCl solution, while DNP is difficult to dissolve in
0.14mol/L NaCl solution. We can get supernate that contain RNP by centrifugation while DNP exist in pellet. Next, Remove protein from RNP to obtain purified RNA. We use chloroform-isoamylol to make protein denature. Therefore protein will form precipitation by centrifugation while RNA stays in supernate. We can get the RNA solution from the liver tissue.
1. Prepare liver homogenate: Take fresh animal liver 30g.Add 200ml 0.14mol/L NaCl solution to prepare 15% liver homogenate with homogeniter.
2. Separate DNP and RNP: Pick up 4ml homogenate to a test tube. Centrifuge
3000rpm/10min.RNP will exist in the supernate while DNP in the precipitation. We transfer the
RNP solution to other test tube.
3. Remove Protein from RNP: Add about 4ml chloroform-isoamylol to the RNP solution.
Churn the tube for about 2min. Centrifuge 3000rpm/10min.The upperest layer is RNA solution.

The medial layer is the denatured protein and the lowest layer is chloroform. We transfer the RNA solution to other tube and discard the rest.
4. Purification of RNA: Add about 6ml 95% ethyl alcohol to the RNA solution. Centrifuge
3000rpm/5min. RNA form precipitation. We should discard the supernate and remain the precipitation. Then add 4ml 0.14mol/L NaCl solution into the tube to dissolve the RNA.
Ⅱ .The Quantification of RNA
The ribose in ribonucleic acid be heated in concentrated acid will dehydrate and become furfurol. Furfurol can react with orcinol agent and become a kind of green compound, which has an absorbance maximum at 670nm.The absorbance is proportional to the concentration of green compound, which is also proportional to the concentration of RNA. So we can calculate the content of RNA in liver tissue by determining the absorbance of the green compound in 670nm .
1. Pick up 3 test tube, add the agents as the table below:
Agents(ml)
RNA standard solution(ml)
RNA sample solution(ml) ddH
2
O(ml) orcinol agent(ml)
Standard tube
1.0
3.0
Sample tube
1.0
3.0
Control tube
1.0
3.0
2.Mix each tube sufficiently, then heat 10 min in the 100 ℃ water bath. Take out the tube and cool to the room temperature. Adjust “0”with the solution in control tube, read the absorption
(A) value of standard and sample tube in 670nm respectively.
3.Calculate:OD sample/OD standard Χ 100 Χ 4/0.5= RNA μ g/g liver tissue
: 100 here is the concentration of RNA standard solution.
4 means that the RNA precipitation is dissolved with 4ml solution.
0.5 means the weight of liver tissue in the experiment.
Ⅲ
See RNA extract. The difference is that DNP can dissolve in 1mol/L NaCl solution while RNP cannot. So we can add 1mol/L NaCl solution to the DNP precipitation to make it dissolve. Then remove protein. It is same to RNA extract.

1. Prepare liver homogenate: See RNA extract.
2. Add about 4ml 1mol/L NaCl solution to the DNP precipitation. Mix the tube sufficiently.
Then lay aside in the room temperature for about 5-6 hours.
3. Remove Protein from DNP: Centrifuge the DNP slurry. Transfer the supernate to other tube and discard the sediment. Add about 4ml chloroform-isoamylol to the tube .Mix the tube sufficiently. Centrifuge 3000rpm/10min.The upperest layer is RNA solution. The medial layer is the denatured protein and the lowest layer is chloroform. We transfer the DNA solution to other tube and discard the rest.
4. Purification of DNA: Add about 6ml 95% ethyl alcohol to the DNA solution. Centrifuge
3000rpm/5min.DNA exist in the precipitation. We should discard the supernate and remain the precipitation. Then add 2ml 1mol/L NaCl solution into the tube to dissolve the DNA.
Ⅳ . The Quantification of DNA
Deoxyribose in DNA be heated in acid solution will dehydrate and become α -hydroxy γ
-ketopentanal. It can react with diphenylamine agent and become a kind of blue compound, which has an absorbance maximum at 595nm.The absorbance is proportional to the concentration of blue compound, which is also proportional to the concentration of DNA. So we can calculate the content of DNA in liver tissue by determining the absorbance of the blue compound in 595nm .
1. Pick up 3 test tube, add the agents as the table below:
Agents(ml)
DNA standard solution(ml)
DNA sample solution(ml) ddH
2
O(ml) diphenylamine agent(ml)
Standard tube
1.0
2.0
Sample tube
1.0
2.0
Control tube
1.0
2.0
2.Mix each tube sufficiently, then heat 10 min in the 100 ℃ water bath. Take out the tube and cool to the room temperature. Adjust “0”with the solution in control tube, read the absorption
(A) value of standard and sample tube in 595nm respectively.
3.Calculate:OD sample/OD standard Χ 400 Χ 2/0.5= DNA μ g/g liver tissue
400 here is the concentration of DNA standard solution.
2 means that the DNA precipitation is dissolved with 2ml solution.

0.5 means the weight of liver tissue in the experiment.
(Liu zhi-fang)

Recombinant DNA technology is genetic engineering, which effects artificial modification of the genetic constitution of a living cell by introduction of foreign DNA through experimental techniques. The technique involves the splicing of DNA by restriction endonucleases, preparation of chimeric DNA molecule by DNA ligase, followed by cloning for the production of identical target DNA molecules.
Most foreign DNA fragments cannot self-replicate in a cell and must therefore be joined to a vector that can replicate autonomously. Each vector typically will join with a single fragment of foreign DNA. If a complex mixture of DNA fragments is used, a population of recombinant DNA molecules is produced. This is then introduced into the host cells, each of which will typically contain only a single type of recombinant DNA. Identification of the cells that contain the DNA fragment of interest allows the purification of large amounts of that single recombinant DNA and hence the foreign DNA fragment.
The following figure shows the typical procedure of recombinant DNA technology:

Tools of recombinant DNA technology: The various “biological tools” used to bring about genetic manipulations include: (1) Enzymes (2) Passenger DNA: foreign DNA (insert DNA fragment) which is passively transferred from one cell to another cell or organ is known as 药店 passenger DNA. (3) Vector or vehicle DNA: The DNA, which acts as the carrier is known as the vector or vehicle DNA.
The various enzymes which may be required to be used are: Restriction endonuclease that cut
DNA chains at specific locations (called as “chemical knife”); Exonucleases that cut DNA at 5′ terminus; Endonuclease that cut in the interior to produce "nicks"; Reverse transcriptase; DNA polymerases; DNA ligase (T4 ligase); S1 nuclease and Alkaline phosphatase.
Passenger DNA (foreign DNA) is DNA to be introduced into the vector DNA. They are genomic DNA fragments, cDNA (complementary DNA), Synthetic DNA, Random DNA.
Vector or Vehicle DNA: Following types of DNA may be used as vector or vehicle DNA:

Bacterial plasmids, Bacteriophages and Cosmids. Bacterial plasmids are of the most importance in our experiments.
Bacterial plasmids are small circular, duplex DNA molecules whose natural function is to confer antibiotic resistance to the host cells. Plasmids DNA replicate independently and they can be easily separated from host bacteria. The DNA sequences and restriction maps of many plasmids are known, hence the precise location of restriction enzyme cleavage sites for inserting the foreign DNA (insert DNA) is available. The plasmids are the most commonly used vectors and can accept short DNA pieces about 6 to 10 kb long.
Plasmid vector pBR322 has both tetracycline (tet) and ampicillin (amp) resistance genes. A single Pst I site. Within the ampicillin resistance gene is commonly used as the insertion site for a piece of foreign DNA. In addition to having sticky ends, the DNA inserted at this site disrupts the ampicillin resistance gene and makes the bacterium carrying this plasmid ampcilllin sensitive.
pUC18/19 belongs to a family of plasmid vectors that contain a polylinker inserted within the alpha region of the lacZ gene. The polylinkers are the same as those used in the M13mp series. pUC18 and pUC19 have the same polylinker but in opposite orientations. Under appropriate conditions, colonies that bear plasmids containing a fragment inserted into the polylinker form white colonies instead of blue ones.
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The ability to isolate, analyze and change genes at will has now become almost commonplace

through recombinant DNA technology. The enormous advances brought about both in understanding gene structure and function and in practical applications of that knowledge depended originally on the development of several major new techniques. Some of the most important were the ability to cut DNA at specific sites using restriction endonucleases, procedures that allow the detection of specific DNA sequences with great accuracy methods for preparing specific DNA sequences in large amounts in pure form and rapid DNA sequencing.
Restriction enzymes are isolated from bacteria, where they play a role in protecting the host cell against virus infection. Over 1800 restriction enzymes have now been isolated and have been named according to the bacterial species from which they were isolated. The first three letters of the enzyme name are the first letter of the genus name and the first two letters of the species name.
Since each bacterium may contain several different restriction enzymes, a roman numeral is also used to identify each enzyme. EcoRI, for example was the first enzyme isolated from Escherichia coli.
Restriction enzymes recognize specific nucleotide sequences in double-stranded DNA that are usually four, five or six nucleotides long, and then cut both strands of the DNA at specific locations. There are basically three ways in which the DNA can be cut: a staggered cut to leave a 5’ overhang, a staggered cut to leave a 3' overhang, or a cut in the same place on both stands to leave a blunt end .For enzymes that cut in the staggered manner, the single strand tails are called cohesive ends because they allow any two DNA fragments produced by the same restriction enzyme to form complementary base pairs .The cut ends can then be joined together by an enzyme called DNA ligase. The new DNA molecule that has been made by joining the DNA fragments is called a recombinant DNA molecule. Blunt-ended DNA molecules can also be joined together by
DNA ligase but the reaction is far less favorable.

When a DNA molecule is cut by a restriction enzyme, the DNA fragments from that restriction digest can be separated by gel electrophoresis. Electrophoresis on a polyacrylamide gel will separate small DNA fragments of less than about 500bp in size, but agarose gels (which have larger pores) are needed to separate larger DNA fragments. The DNA digest separates into a series of bands representing the restriction fragments. Since small fragments travel further in the gel than larger fragments, the size of each fragment can be determined by measuring its migration distance relative to standard DNA fragments of known size. The DNA can be located after gel electrophoresis by staining with ethidium bromide that binds to the DNA and fluoresces a bright orange. Alternatively, if the DNA is labeled with a radioisotope such as 32 P, the bands can be detected after electrophoresis by laying the gel against an X-ray film whereby the radioactivity causes silver grain to be formed in the film emulsion, giving black images corresponding to the radioactive bands.
Alkaline lysis, in combination with the detergent SDS, has been used for more than 20 years to isolate plasmid DNA from E.coli. Exposure of bacterial suspensions to the strongly anionic detergent at high pH opens the cell wall, denatures chromosomal DNA and proteins, and releases plasmid DNA into the supernatant. Although the alkaline solution completely disrupts base pairing, the strands of closed circular plasmid DNA are unable to separate from each other because they are topologically intertwined. As long as the intensity and duration of exposure to OH - is not too great, the two strands of plasmid DNA fall once again into register when the pH is returned to neutral. Alkaline lysis in the presence of SDS is a flexible technique that works well with all strains of E.coli and with bacterial cultures ranging in size from l ml to >500ml.The closed circular plasmid DNA recovered from the lysate can be purified in many different ways and to different extents, according to the needs of the experiment.
During lysis, bacterial proteins, broken cell walls, and denatured chromosomal DNA become enmeshed in large complexes that are coated with dodecyl sulfate. These complexes are efficiently precipitated from solution when sodium ions are replaced by potassium ions. After the denatured

material removed by centrifugation, native plasmid DNA can be recovered from the supernatant.
Agarose is a linear polymer composed of alternating residues of D- and L-galactose joined by
(1→3) and (1→4) glycosidic linkages. The L-galactose residue has an anhydro bridge between the three and six positions. Chains of agarose form helical fibers that aggregate into supercoiled structures with a radius of 20-30nm.Gelation of agarose results in a three-dimensional mesh of channels whose diameters range from 50nm to 200 nm.
Commercially prepared agarose polymers are believed to contain ~800galactose residues per chain. However, agarose is not homogeneous: The average length of the polysaccharide chains varies from batch to batch and from manufacturer to manufacturer. In addition, lower grades of agarose may be contaminated with other polysaccharides, as well as salts and proteins. This variability can affect the gelling/melting temperature of agarose solutions, the sieving of DNA, and the ability of the DNA recovered from the gel to serve as a substrate in enzymatic reactions.
These potential problems can be minimized by using special grades of agarose that are screened for the presence of inhibitors and nucleases and for minimal background fluorescence after staining with ethidium bromide.
The following factors determine the rate of migration of DNA through agarose gels:
1.)The molecular size of the DNA. Molecules of double-stranded DNA migrate through gel matrices at rates that are inversely proportional to the log
10 of the number of base pairs. Larger molecules migrate more slowly because of greater frictional drag and because they worm their way through the pores of the gel less efficiently than smaller molecules.
2.)The concentration of agarose. A linear DNA fragment of a given size migrates at different rates through gels containing different concentrations of agarose. There is a linear relationship between the logarithm of the electrophoretic mobility of the DNA (μ) and the gel concentration (l) that is described by the equation: log μ =log μ。一 K r l where μ。 is the free electrophoretic mobility of DNA and K r
is the retardation coefficient, a constant related to the property of the gel and the size and shape of the migrating molecules.
3.)The conformation of the DNA. Superhelical circular (form I), nicked circular (form II), and

linear (form III) DNAs migrate through agarose gels at different rates. The relative mobilities of the three forms depend primarily on the concentration and type of agarose used to make the gel, but they are also influenced by the strength of the applied current, the ionic intensity(strength) of the buffer and the density of superhelical twists in the form I DNA. Under some conditions, form I
DNA migrates faster than form III DNA; under other conditions, the order is reversed. In most cases, the best way to distinguish between the different conformational forms of DNA is simply to include in the gel a sample of untreated circular DNA and a sample of the same DNA that has been linear zed by digestion with a restriction enzyme that cleaves the DNA in only one place.
4.)The presence of ethidium bromide in the gel and electrophoresis buffer. Intercalation of ethidium bromide causes a decrease in the negative charge of the double-stranded DNA and an increase in both its stiffness and length. The rate of migration of the linear DNA-dye complex through gels is consequently retarded by a factor of ~15%.
5.)The applied voltage. At low voltages, the rate of migration of linear DNA fragments is proportional to the voltage applied. However, as the (strength) of the electric field is raised, the mobility of high-molecular-weight fragments increases differentially. Thus, the effective range of separation in agarose gels decreases as the voltage is increased. To obtain maximum resolution of
DNA fragments >2kb in size, agarose gels should be run at no more than 5-8V/cm.
6.)The type of agarose. The two major classes of agarose are standard agaroses and low-melting-temperature agaroses. A third and growing class consists of intermediate melting/gelling temperature agaroses, exhibiting properties of each of the two major classes.
7.)The electrophoresis buffer. The electrophoretic mobility of DNA is affected by the composition and ionic strength of the electrophoresis buffer. In the absence of ions, electrical conductivity is minimal and DNA migrates slowly, if at all. In buffer of high ionic strength (e.g if 10 × electrophoresis buffer is mistakenly used ),electrical conductance is very efficient and significant amounts of heat are generated, even when moderate voltages are applied. In the worst case, the gel melts and the DNA denatures.
Gel-loading buffers are mixed with the samples before loading into the slots of the gel. These buffers serve three purposes: They increase the density of the sample, ensuring that the DNA sinks evenly into the well; they add color to the sample, thereby simplifying the loading process; and they contain dyes that, in an electric add, move toward the anode at predictable rates.
Bromophenol blue migrates through agarose gels ~2.2-fold faster than xylene cyanol FF, independent of the agarose concentration. Bromophenol blue migrates through agarose gels run in
0.5x TBE at approximately the same rate as linear double-stranded DNA 300bp in length, whereas xylene cyanol FF migrates at approximately the same rate as linear double-stranded DNA 4kb in length. These relationships are not significantly affected by the concentration of agarose in the gel over the range of 0.5-1.4%. Which type of loading dye to use is a matter of personal preference.
The most convenient and commonly used method to visualize DNA in agarose gels is staining with the fluorescent dye ethidium bromide, which contains a tricyclic planar group that intercalates between the stacked bases of DNA. Ethidium bromide binds to DNA with little or no sequence preference. At saturation in solutions of high ionic strength, approximately one ethidium molecule is intercalated per 2.5bp.After insertion into the helix, the dye lies perpendicular to the helical axis and makes vander Waals contacts with the base pairs above and below. The axed position of the planar group and its close proximity to the bases cause dye bound to DNA to display an increased fluorescent yield compared to that of dye in free solution. UV radiation at

254nm is absorbed by the DNA and transmitted to the dye; radiation at 302nm and 366nm is absorbed by the bound dye itself; In both cases, the energy is re-emitted at 590 nm in the red-orange region of the visible spectrum. Because the fluorescent yield of ethidium bromide-DNA complexes is 20~30-fold greater than that of unbound dye, bands containing as little as 10ng of DNA can be detected in the presence of free ethidium bromide (0.5μg/ml) in the gel.
Agarose gel electrophoresis of DNA fragment. DNA fragments of known size were electrophoresed in lane 1(the sizes in bp are given on the left). A restriction digest of the sample
DNA was electrophoresed in lane 2. By comparison with the migration positions of fragments in lane 1, it can be seen that the two sample DNA fragments have sizes of approximately 9000 and
2000 bp. The sizes could be determined more accurately by plotting the data from lane 1 as a standard curve of log DNA size vs migration distance and then using this to estimate the size of the sample DNA fragments from their measured migration distances.
1.Transfer a single bacterial colony into 3.5 ml of LB liquid medium containing of the appropriate antibiotic (Amp 60μg/ml LB) in a loosely capped tube. Incubate the culture overnight at 37 ° C with vigorous shaking.
2.Pour 1.5ml of the culture into a microfuge tube (1.5ml). Centrifuge at 8,000rpm for 30 seconds in the microfuge. This step could be repeated for acquiring more bacteria.
3.Remove the medium by pouring out, leaving the bacterial pellet as dry as possible.
4.Resuspend the bacterial pellet obtained in step 3 above in 100 μl of ice-cold Solution Ⅰ by vigorous vortex.

5.Add 200μl of freshly prepared Solution Ⅱ . Close the tube tightly, and mix the contents by inverting the tube rapidly five to ten times. Do not vortex. Store the tube for 3-5 minutes at room temperature.
6.Add 150 μl of solution Ⅲ .Close the tube and vortex it gently in an inverted position for 10 seconds to disperse Solution Ⅲ through the viscous bacterial lysate. Store the tube for 3-5 minutes at room temperature. Centrifuge at 10,000rpm for 5 minutes in a microfuge.
7.Transfer the supernatant to a fresh microfuge tube. Add an equal volume of chloroform. Mix by vortex. After centrifuging at 5,000rpm for 30 seconds in a microfuge, transfer 80% of the supernatant to a fresh tube.
8.Precipitate the double-stranded DNA with 2 volumes of ethanol at room temperature. Mix by vortex. Allow the mixture to stand for 5 minutes at room temperature. Centrifuge at 10,000rpm for
5 minutes in a microfuge.
9.Discard the supernatant and rinse the pellet of double-stranded DNA with 1ml of 70% ethanol.
Remove the supernatant as possible as you can, and allow the pellet of nucleic acid to dry in the air for 10 minutes.
10.Redissolve the nucleic acid in 40μl of TE containing DNAase-free pancreatic RNAase
(20μg/ml). Vortex briefly.
The sample of plasmid DNA could be identified by agarose electrophoresis.
Ⅲ
Ⅰ
1.Add 5ul of 10 × H buffer, 3ul of ddH
2
O and 2ul of Hind Ⅲ to 40ul plasmid solutions
(containing pUC18/19 or pBR322), the ultimate volume is 50ul.
2.Incubate for 2 hours at 37 ℃
3.Digesting with enzyme Hind Ⅲ ,and then take 10ul of the sample to be identified by agarose electrophoresis.
4.The residual sample (40ul) then is digested with enzyme Pst Ⅰ :
5.Add 4ul of 3mol/L NaAc to the sample above, mix by vortex
6.Add 2 volumes of ethanol (88 ul) at room temperature. Mix by vortex. Allow the mixture to stand for 5 minutes at room temperature. Centrifuge at 10,000rpm for 5 minutes in a microfuge.
7.Discard the supernatant and rinse the pellet of double-stranded DNA with 1ml of 70% ethanol.
8.Remove the supernatant, and allow the pellet of nucleic acid to dry in the air for 10 minutes.
9.Redissolve the nucleic acid in 30μl of TE. Vortex briefly.
10.Add 4ul of 10 ×( H ?)buffer, 2ul of PstI and 4ul of ddH
2
O to 30ul plasmid solutions
(containing plasmid pUC18/19 or pBR322 digested by Hind Ⅲ ), the ultimate volume is 40ul.
11.Incubate for 2 hours at 37 ℃ , then take 10ul of the sample out to be identified by agarose electrophoresis
.
Seal the edges of clean, dry open ends of the plastic plate with autoclave adhesive tape to form a mold. Set the mold on a horizontal section of the desk. Position the comb 0.5-1.0mm above the

plate so that a complete well is formed when the agarose is added. If the comb is closer to the plate, there is a risk that the base of the well may tear when the comb is withdrawn, allowing the sample to leak between the gel and the plate.
1.
Pour the warm agarose solution into the mold. The gel should be 3mm to 5mm thick. Check to see that there are no air bubbles under or between the teeth of the comb.
2.
After the gel is completely set (30-45 minutes at room temperature), carefully remove the comb and the tape and mount the gel in the electrophoresis tank. Add just enough buffer to cover the gel to a depth of about 1mm.
3.
Mix the sample of DNA with the desired gel-loading buffer. Slowly load the mixture into the slots of the submerged gel using a disposable micropipette.
4.
Close the lid of the gel tank and attach the electrical leads so that DNA will migrate toward the anode, if the leads have been attached correctly, bubbles should be generated at the anode and cathode and within a few minutes the bromophenol blue should migrate from the wells into the body of the gel. Run the gel until the bromophenol blue migrates the appropriate distance through the gel. Turn off the electric current and remove the leads and lid from the gel tank.
5.
Examine the gel by ethidium bromide and photograph the gel as described.
(Guo Qiang)