Preface Science and technology progressed by leaps and bounds in this century, particularly in the field of Biotechnology and Molecular Biology. Keeping pace with these advances UGC has restructured its curricula and has introduced new experiments at undergraduate and postgraduate levels in various universities and professional courses. Every University and institutions have introduced “Biotechnology” in their syllabus, either as compulsory paper or as vocational paper. It has thus become relevant that infrastructure facilities in life science should be provided in different colleges and universities so that our students should be able to compete nationally and internationally. Yet, for reasons not known there lies a wide range of disparity between the theoretical and practical studies. As a result both teachers and students of undergraduate and post graduate level face tremendous difficulties in acquiring study material of practical utility, particularly topics related to DNA Isolation, Plasmid Isolation, Competent cell preparation, Cloning experiments, PCR, SDS-PAGE, separation techniques for DNA, RNA and proteins. After the success of our previous book “Experimental Biology-A Laboratory Manual”, which presents both the principles of analytical biology that makes experimental, analytical and separation techniques possible in the laboratory, with background information to understand what the students are going to do and why, including technical information the students gather in the laboratory, we thought it would be logical to start our new venture on “Experimental Biotechnology-A Laboratory Manual”. The book is subdivided into seven sections This encompass: General procedures, like methods of pipetting, solution preparation, buffers and principles of common analytical instruments essential for laboratory biotechnology experiments. The book also includes working with nucleic acid, bacteria, enzymes, proteins; Cloning experiments and a few protocols on Plant biotechnology. Emphasis have been given on DNA/RNA isolation from various sources, use of restriction enzymes, ligation techniques, cloning protocols, screening of transformed cells, various electrophoresis techniques, PCR protocol, etc. The appendices in the last part are included to provide information important to the study of the above mentioned practical as a whole.

This book is designed to introduce to the students some of the most widely used experimental procedures in biotechnology, including DNA/RNA isolation, manipulation, cloning, protein purification and characterization. The students will also get familiarized with some of the equipments frequently used in biochemistry and molecular biology. The biotechnology laboratory course is an exploration of procedures. Hence, in order to get full benefit from the course, the students need to read the manual, which is self explanatory. The more effort they put into the course work, the more they will learn. Before the start of actual lab works, one needs to spend some time reading the Laboratory Manual. This reading will provide background information and an outline of the procedures to be performed. The biotechnology laboratory is conducted as a “directed” research project. This means that although the general procedures are well established, the overall goal of each experiment is the acquisition of new information. Because of the nature of scientific research, predicting the outcome of experiments that have not previously been performed is difficult. It may therefore be necessary to design new experiments based on the results of previous ones, or to repeat experiments that yielded uninterruptable or ambiguous results.

In molecular biology and biochemistry, the ability to accurately and reproducibly measure and transfer small volumes of liquids is critical for obtaining useful results. For volumes less than 1 ml, the most common method for measuring liquid volumes involves the use of a device known as a pipetman or micro-pipette. The pipetmen can be both fixed and variable volumes. Commonly used pipetmen are: P1000, P200, P20, P10 etc. P1000 are useful for volumes from 200 to 1000 µl. P200 are useful for volumes from 20 to 200 µl. P20 are useful for volumes from 0.5 to 20 µl. For beginners fixed volume pipetmen are more useful. It is important to ascertain whether the correct pipetman is chosen for measuring the volume one needs.

A spectrophotometer is an analytical instrument that measures the absorbance of a test sample (solution). The term “spectroscopy” comes from the word “spectrum”. “Spectroscopy” therefore implies the use of multiple wavelengths of light. Spectrophotometers have the ability to specifically measure absorbance at specific wavelengths. The most commonly used method to allow this involves a “monochromator”, a device (either a prism, or more commonly, a diffraction grating) that splits the incident light into its component wavelengths, and allows only light of the desired wavelength to reach the sample. The ability to measure absorbance at different wavelengths is very useful, because the extinction coefficient of a compound varies with wavelength. In addition, the absorbance spectrum of a compound can vary depending on the chemical composition of the compound, and depending on the nature of the solvent around the compound. Absorbance is a useful quantity and follows the Beer-Lambert law. The extinction coefficient of a molecule at a given wavelength can be calculated using the Beer-Lambert equation from absorbance measurements for solutions of known concentration, which states that:

To prepare a series of dilutions of the original more concentrated stock solution, one need to consider both the required final concentration and required volume of the diluted material. The dilution can be readily calculated by the equation:

Several substrates, like proteins (enzymes), are very sensitive to alteration in the concentration of the solution they are placed in. Hence is the importance of a buffer which controls the chemical properties of the medium. Literally a “buffer” has the ability to resist changes in the hydrogen ion concentration. Although, buffers do contain other molecules, that might, in addition to the hydrogen ion concentration, influence the ionic strength, the activity of enzymes, and other parameters of the experiment. In any biological experiment, appropriate buffer must be selected based on their effect on the experiment. Ideal buffers control pH and ionic strength without interacting adversely with the experiment undertaken. For example, although phosphate is a common buffer, it may not be suited when phosphate is a substrate or product of the reaction being studied. Also, some proteins interact poorly with some buffers. For example, Tris is not ideal as it exhibit high pKa and the large change in pKa when temperature changes, although it is cheap and most proteins are stable in Tris buffers, moreover it seldom reacts with reaction substrates hence, Tris buffer is commonly used in biochemical experiments.

Autoclave Operating Procedures Place all material to be autoclaved in an autoclavable tray. All items should have indicator tape. Separate liquids and solids should be autoclaved separately. Ensure that the lids on all bottles are loose. Do not put a large number of items in tray, to ensure that all items reach the appropriate temperature. Also allow sufficient air/steam circulation. Steps in autoclave Pressure should read ‘O’ when you open the pressure chamber of the autoclave. Place items to be autoclaved in the autoclave and close the door. Some autoclaves require that you also lock the door after it’s closed. Set time - typically 20 minutes. Temperature should be set at 121°C already, but double-check and change if necessary. Set cycle: If liquid, set “liquid cycle” or “slow exhaust”. If dry, set “dry cycle” or “fast exhaust” + dry time. Start the cycle. On some autoclaves, the cycle starts automatically at step 5. On others, turn to “sterilize”

Storage Nucleic acids are very sensitive to contaminations hence care should be taken while processing and storing DNA and RNA. The following properties of reagents and conditions need to be taken care of: Heavy metals cause breakage of phosphodiester linkage. One can use EDTA which is an excellent heavy metal chelator. UV rays cause extensive damage to nucleic acids. UV light at 260 nm causes breakage of thymine dimers and cross-link, including loss of biological activities. Ethidium bromide results in photo oxidation of DNA. Oxidation products can cause phosphodiester breakage. Nucleases are found on human skin; therefore, it is advisable to avoid direct or indirect contact between nucleic acids and fingers. Most DNAes are not very stable; however, many RNAses are very stable and can absorb to glass or plastic and remain active. At -20°C: extensive single and double strand breaks results. Hence for long-term storage it should be stores at -70°C. ? It is best to store DNA in high salt ( >1M) in the presence of high EDTA (>10mM) at pH 8.5, for long-term storage. Storage of DNA in CsCl with ethidium bromide in the dark is an excellent storage medium. There is about one phosphodiester break per 200 kb of DNA per year. Storage of ë DNA in the phage is better than storing the pure DNA.

Cell Genotypes A large variety of bacterial strains, like E. coli, are frequently used in molecular biological experiments. Students should choose the strain or strains required for an experiment on the basis of genetic properties of the strains. It is thus pertinent that a sound knowledge of the genetic makeup of the E. coli stains is acquired. The genetic makeup of the strains of E.coli is distinctly different from the wild type (K12 or B strains) strains from which they are derived and contains genes that can be modified. Genes are supposed to be mutated when they are not listed in a genotype. In most cases, the genes listed are mutated to the point of inactivity. In a few cases, a “-” (indicating inactive or absent or deleted), “+” (indicating active). , The genotype is a factor in choosing a bacterial strain. Chosen strains should be ascertained for their utility for the specific experiment. Small Scale Cultures Fresh culture from a freshly streaked plate should be taken while experimenting with E. coli cells. For preparing a small scale culture of E. coli , prepare 3-5 ml of LB medium/broth (or appropriate broth include antibiotic if the culture contains a plasmid) in two sterile 50 ml tubes. Inoculate one tube with a single colony from a fresh plate. The second tube is used as a broth control. Incubate both tubes at 37°C, shaking vigorously overnight. Inspect the tubes the next morning. The broth control should be clear and the inoculated culture should be very turbid. Make a note of any debris found in the tubes and only incubate longer if the culture is not dense. Never allow cells to overgrow. For some applications, cells can be stored at 4°C for short periods prior to use.

PCR Principles and Procedure Polymerase chain reaction (PCR) is a technique that allows the generation of large amounts of a single DNA sequence from a mixture of sequences; the fragment generated can be designed to contain specific starting and ending positions based on the needs of the experiment. PCR is used to amplify specific regions of a DNA strand (the DNA target). This can be a single gene, a part of a gene, or a non-coding sequence. Most PCR methods typically amplify DNA fragments of up to 10 kilo base pairs (kb), although some techniques allow for amplification of fragments up to 40 kb in size. A basic PCR set up requires several components and reagents. These components include:

Principle Apart from PCR for generating large amounts of DNA, the second method is to use bacteria to replicate the DNA and then purify the replicated DNA from the bacteria. Plasmid DNA is usually the DNA of interest for this method. A plasmid is a double stranded circular DNA molecule that will replicate in an organism. A typical plasmid contains four important features: The plasmid must be circular, because bacteria generally will not replicate linear ? DNA. The plasmid must contain a sequence that functions as an origin of replication (ori). The plasmid must contain a selection mechanism that will force the bacteria to retain the DNA; the most common type of selection mechanism used in bacteria is a gene for resistance to an antibiotic such as ampicillin or tetracycline. The plasmid must contain a region for the insertion of the experimental DNA. A generic plasmid exhibiting these features is shown at right. An expression plasmid that allows expression of foreign DNA should have a strong promoter segment for transcription of inserted DNA. Must have an effective ribosome binding site that will allow efficient translation of the transcribed RNA. Must have high copy number, i.e., the host bacterium should have many copies (usually >100)

Isolation of plasmid DNA from E. coli is a common routine in research laboratories. One can perform a widely-practiced procedure that involves alkaline lysis of cells. This protocol often referred to as a plasmid “mini-prep,” yields fairly clean DNA quickly and easily. Procedure Fill a microcentrifuge tube with saturated bacterial culture grown in LB broth + antibiotic. Spin tube in microcentrifuge for 1 minute, and make sure tubes are balanced in microcentrifuge. Dump supernatant and drain tube briefly on paper towel. Repeat step 1 in the same tube, filling the tube again with more bacterial culture. The purpose of this step is to increase the starting volume of cells so that more plasmid DNA can be isolated per prep. Spin tube in microcentrifuge for 1 minute. Pour off supernatant and drain tube on paper towel. Add 0.2 ml ice-cold Solution 1 to cell pellet and resuspend cells as much as possible using disposable transfer pipet.

Principle The principle is to extract plasmid DNA from bacterial cell suspensions and is based on the alkaline lysis procedure developed by Birnboim and Doly (Nucleic Acids Research 7:1513, 1979). It is a known fact that plasmids are much smaller super coiled DNA molecules than the much larger and less super coiled bacterial chromosomal DNA. This difference in topology enables selective precipitation of the chromosomal DNA and cellular proteins and separates them from plasmids and RNA molecules. The cells are lysed under alkaline conditions, which denature both nucleic acids and proteins, and when the solution is neutralized by the addition of Potassium Acetate, chromosomal DNA and proteins precipitate because it is impossible for them to renature correctly (they are so large). Plasmids renature correctly and stay in solution, effectively separating them from chromosomal DNA and proteins.

Solution Preparation 1. Glycerol mix (1 L) Weigh 25 grams (liquid weight) of glycerol and Add dH2O to 1 Liter (Autoclave). 2. Potassium mix (1 L) 170 mL of 1M KH2PO4 (monobasic), 720 mL of 1M K2HPO4 (dibasic), and 110 mL of dH2O (Autoclave)

Preparation of Solutions GTE buffer 50 mM Glucose, 25 mM Tris-Cl, 10 mM EDTA, pH 8 NaOH/SDS lysis solution 0.2 M NaOH, 1% SDS

Transfer 1.5 mL overnight culture to a 1.5 mL microfuge tube, centrifuge for 30 sec, and decant supernatant. Resuspend cells in 400 µL TE by vortexing, add 50 µL 10% SDS, 50 µL proteinase K (20 mg/mL in TE). Incubate for 1 hour at 37oC. Shear DNA by 3-5 passages through a 26 G needle. Extract twice with 500 µL phenol: chloroform (1:1), and twice with 500 µL chloroform.  Precipitate nucleic acids by adding 25 µL 5 M NaCl and 1 mL 95% EtOH, vortex, and centrifuge for 10 min, decant supernatant. Dry pellet.

Reagents EDTA Ethanol, absolute Isopropanol Phosphate Buffered Saline (PBS), 1X Proteinase K Sodium Acetate, 3 M pH 5.2 (Molecular Biology Grade) Sodium chloride (NaCl) Sodium Dodecyl Sulfate (SDS ), 10% Tris EDTA (TE), pH 7.4 Tris EDTA (TE), pH 8.0 Tris HCl, pH 8.0

Reagents Chloroform EDTA, 0.5 M Ethanol, absolute Isoamyl alcohol Phenol Phosphate Buffered Saline (PBS), 1X Proteinase K RNAse A Sodium dodecyl sulfate (SDS) solution, 10%

Reagents Chloroform EDTA, 0.5 M Ethanol, absolute Is amyl alcohol Phenol Proteinase K RNase A Sodium acetate, pH 5.2 Sodium chloride, 5 M Sodium thiocyanate (NaSCN), 1 M TE Buffer (Tris-EDTA), pH 7.4 Tween 20 Xylene

Principle Restriction digestion is one of the most common reactions performed in molecular biology. Restriction endonucleases (RE) are also known as restriction enzymes, which recognize short, specific DNA sequences. There are three types of RE: Type-I, Type-II and Type-III. Type II RE, most commonly used for DNA analysis and genetic engineering, cleave double-stranded DNA (dsDNA) at specific sites within or adjacent to their recognition sequences. All Type II RE recognises a unique nucleotide sequence at which it cuts a DNA molecule and nowhere else. The recognition sequence is often a six base pair palindromic sequence (the top DNA strand from 5' to 3' is the same as the bottom DNA strand from 5' to 3'), but others recognize four or even eight base pair sequences. Many RE will not cut DNA that is methylated on one or both strands of the recognition site, although some require substrate methylation. REs can also differ in the way they cut the DNA molecule. Some enzymes cut in the middle of the recognition sequence, resulting in a flush or “blunt end”. Other enzymes cleave in a staggered fashion, resulting in DNA products that have short single-stranded overhangs (usually two or four nucleotides) at each end. These are often called “cohesive ends”, as these single-stranded overhangs could potentially come together again through complementary base-pairing, as shown below:

Principle Since the entire DNA sequence of the bacteriophage is known, we know exactly where a restriction enzyme will cleave the viral DNA and the size of the DNA fragments generated. This restriction enzyme digest will create a pattern of specific DNA fragments that will serves as a “DNA ladder”.

Reagents Nuclease-Free Water: 14µl 10X Restriction Buffer: 2µl Acetylated BSA (1mg/ml): 2µl DNA (~1µg): 1µl Restriction Enzyme (10u): 1µl Final Volume: 20µl

Reagents Restriction enzymes of choice, such as BamH1 and EcoRI Restriction enzyme reaction buffer, 70 % Ethanol 100 % Ethanol 3 M Sodium acetate (pH 5.2) Distilled water Plasmid DNA isolated from bacteria, such as pGFP(R) and pBC(R)

Principle of DNA Ligase Mechanism The process of connecting two pieces of DNA together is called ligation, and is catalyzed by an enzyme called ligase. The ligase used most often in molecular biology is derived from the T4 bacteriophage, and uses ATP to supply the energy necessary for the reaction. In addition to ATP, T4 ligase requires DNA with a 5´-phosphate group and free 3´-hydroxyl groups. Applications Ligation of blunt or cohesive-ended DNA fragments. Repair of nicks in double-stranded nucleic acids.

Start with a 1:2 molar ratio of vector: insert DNA when cloning a fragment into a plasmid vector. This ratio will vary with other types of vectors, for example, cDNA and genomic cloning vectors. The following example illustrates the conversion of molar ratios to mass ratios for a 3.0kb plasmid and a 0.5kb insert DNA fragment.

Principle The rate of joining of DNA fragments is affected by seven parameters: DNA concentration, Concentration of compatible ends, Size of the fragments, Presence or absence of complementary, single-stranded ends (some nucleases form blunt ends and in these cases special techniques are needed for joining), G + C content and length of the complementary single strands, Temperature of the reaction and Ionic composition of the solution.

Principle Plasmids requires hosts to replicate, hence it has to be inserted into bacteria. But efficiency of uptake of plasmid DNA by bacteria is low and will not do so without getting degraded. Therefore, to enhance the probability of uptake of the plasmid DNA by bacteria it is necessary to make the bacterial cells “competent” to absorb the plasmid DNA. One of the methods to increase efficiency of cell competence is given below. It should be noted that competent cells are more fragile than normal bacteria. Vortexing the cells, heating the cells above 42ºC or to 42°C for prolonged periods, or exposure of the cells to abusive treatments may kill them

Steps for Cloning Prepare DNA. Select restriction enzymes for your insert and vector, and determine the appropriate reaction buffers. Combine the following in a microfuge tube (30 µL total volume): 2 µg DNA 1 µL Each Restriction Enzyme 3 µL 10x Buffer 3 µL 10x BSA x µL H2O (to bring total volume to 30 µL)

Transformation The uptake of DNA by competent cells is called transformation. Transformation involves: Mix a small volume of DNA (usually 1-3 µl) with 100 µl of competent cells. The mixture is incubated on ice for 30 minutes to allow the cells to take up the DNA. The cells are then heat-shocked at 37 to 42°C for 1 minute, and cold shocked on ice for 2 minutes. The heat shock/cold shock reverses the effect of the competent cell process, by healing the damage inflicted by the competent cell solution. Add 0.5 ml of LB culture mediumz Incubate the cells for 10 - 30 minutes at 37°C to aid the healing process. This cell/medium mixture is then spread onto bacterial culture plates containing agar, LB, and a selection mechanism.

Principle Cloning experiments would be very difficult if we do not screen those very few cells that took up the recombinant plasmid DNA, as the transformation efficiency is very low as stated before (usually less than 1 in 105 cells). Hence it is necessary that there should be a selection mechanism (i.e. conditions where cells grow if they express a gene, but die if they do not) to sort out which are transformed cells and which are not make the job easy. A commonly used selection mechanism is antibiotic resistance. If the E. coli strain used is not resistant to the antibiotic, but the plasmid DNA contains a gene coding for resistance, then only the cells that have taken up the plasmid will be able to grow. Most commonly used plasmids contain a gene for resistance to ampicillin; only cells that have taken up the plasmid will be able to grow and form colonies. Note: Uptake of plasmid DNA is rare; hence each group of cells visible on a plate represents the offspring of a single cell that will contain a single molecule of plasmid DNA.

Principle The principle of Electrophoresis involves the motion of a particle in a fluid medium, where the gravitational attraction (on a flowing liquid) is the sedimenting force, and the frictional force is the opposing force and is proportional to the velocity of the particle. These forces balance each other acting in opposite directions and hence balance each other. This will lead to the uniform motion of the particle in the mobile liquid medium (i.e. the particle moves at constant velocity). In case the external field is an electric field instead of a gravitational field, a macromolecule will respond to the external field in two ways. In case the molecule is charged, it will migrate in an electric field to the electrode of opposite charge. Under this principle the technique of electrophoresis functions. All electrophoresis operations are governed by the principle:

Agarose gel electrophoresis is the easiest and commonest way of separating, capable of resolving mixtures of DNA fragment that cannot be separated conveniently by other procedures. The purpose of the gel might be to look at the DNA, to quantify it or to isolate a particular band. The DNA is visualised in the gel by addition of ethidium bromide. This binds strongly to DNA by intercalating between the bases and is fluorescent meaning that it absorbs invisible UV light and transmits the energy as visible orange light. Agarose, which is extracted from seaweed, is a linear polymer whose basic structure is shown in figure.

here are many methods for extracting DNA from gel apart from using commercial kits, some methods however yield cleaner DNA than others. 1. Freeze squeeze Excise band and place pieces of it in a home-made spin-column (a piece of filter placed in a PCR 0.5ml eppendorf, pierced the bottom of the PCR tube with a needle). Place the PCR tube at -20 °C for 15-20 mins or freeze in liquid N2. Then place the PCR tube into the 1.5 ml eppendorf and spin at top speed in a microfuge for 15 mins, collect DNA solution which can be EtOH ppt or use directly. 2. Powdered silica Melt excised bands at 70°C cool and add equal volume of TE-buffered phenol and mix. Spin for 5 min, remove aqueous phase, repeat and EtOh ppt. the DNA. 3. Electroelution

Principle To retrieve and purify any specific DNA fragment from an agarose gel slice; the expected yield is from 50-75% of the amount in the gel slice. Time required Restriction digest - minimum of 2 hours Electrophoresis - approx. 4 hours Elution - 2-5 hours depending on fragment size DNA Purification -2 hours

Principle RNA is negatively charged at neutral pH and when an electric field is applied, it migrates towards the anode. RNA retains much of its secondary structure during electrophoresis unless it is first denatured. The addition of formaldehyde to the agarose gel maintains the RNA in its linear (denatured) form. The following protocol is based on Formaldehyde Agarose Gel Method.

Principle Monomeric acrylamide (which is neurotoxic) is polymerised in the presence of free radicals to form polyacrylamide. The free radicles are provided by ammonium persulphate and stabilised by TEMED (N’N’N’N’-tetramethylethylene-diamine). The chains of polyacrylamide are cross-linked by the addition of methylenebisacrylamide to form a gel whose porosity is determined by the length of chains and the degree of crosslinking. The chain length is proportional to the acrylamide concentration: usually between 3.5 and 20%. Cross-linking bisacrylamide is usually added at a ratio of 2g BIS 38g acrylamide.

Denaturing polyacrylamide/urea gels in TBE buffer Prepare 20ml of a 5% polyacrylamide gel containing 7M urea by adding: 47.5% acrylamide: 2.5% bis-acrylamide solution 2ml 10M urea 14ml 10X TBE buffer 2ml 10% freshly prepared ammonium persulfate 0.2ml deionized water 1.8ml

The principle of silver nitrate staining of proteins is based on the reduction of silver ions to its metallic form. (Silver ions in alkaline conditions bind with proteins through å-amino group of lysine and sulphur groups of cysteine and methionine residues). When the complexed silver ions are reduced in the presence of formaldehyde, they become metallic silver, which is seen on the gel as bands. Staining of proteins with silver nitrate after SDS-PAGE is quicker and ~ 10 times more sensitive than Coomassie brilliant blue staining and that is the reason it is widely used in research laboratories these days. The procedure is as follows:

Principle Determination of the precise sequence of nucleotides in a sample of DNA is called “DNA Sequencing”. The most popular method for doing this is called the “Dideoxy method or Sanger Method” (named after Frederick Sanger, who was awarded the 1980 Nobel Prize in chemistry for inventing it). DNA is synthesized from four deoxynucleotide triphosphates. The top most figure is a normal nucleotide: deoxythymidine triphosphate (dTTP). During polymerization each new nucleotide is added to the 3' -OH group of the last nucleotide added. The dideoxy method has been coined after the synthetic nucleotides that lack the –OH at the 3’carbon atom of the sugar and it plays an important role in this process atom (red arrow). In the figure below dideoxythymidine triphosphate — ddTTP, a dideoxynucleotide has been shown that can be added to the growing DNA strand. But the chain elongation ceases when the next nucleotide tries to get attached to the former as there is no 3' -OH for the next nucleotide to be attached to. Hence, the dideoxy method is also called the chain termination method. 

Principle The SDS polyacrylamide gel electrophoresis (PAGE) performs: Separation of proteins according to their molecular weight Ascertain whether a given protein is actually present in a sample, Assessment of purity of the preparation Estimation of the approximate quantity of the protein, and Measurement of the size of the protein.

Cell Lysis Procedure Pellet the cells in a microfuge tube: add 1.5 ml culture, centrifuge for 15 seconds at maximum speed, discard the supernatant, and then repeat the procedure. This will result in the cells from 3 ml of culture in a single tube. Resuspend the cells in 1 ml Lysis buffer. Take 300 µl aliquot for Bradford protein assay. Add 50 µl lysozyme, and mix gently. After 10-20 minutes at room temperature, microfuge for 10 minutes to pellet cell debris. Use supernatant for LDH assay.

Principle Western blot analysis (also called protein immunoblot) is a useful tool for the detection of proteins in biological samples. It uses gel electrophoresis to separate native or denatured proteins by the length of the polypeptide (denaturing conditions) or by the 3-D structure of the protein (native/ non-denaturing conditions). The proteins are then transferred to a membrane (typically nitrocellulose), where they are probed (detected) using antibodies specific to the target protein.  It is a technique for analyzing and identifying protein antigens: the proteins are separated by electrophoresis in polyacrylamide gel, then transferred (“blotted”) onto a nitrocellulose membrane or treated paper, where they bind in the same pattern as they formed in the gel. The antigen is overlaid first with antibody, then with anti-immunoglobulin or protein A labeled with radioisotope, fluorescent dye, or enzyme.  The method originated from the laboratory of George Stark at Stanford. The name “Western blot” was given to the technique by W. Neal Burnette and is a play on the name “Southern blot”, a technique for DNA detection developed earlier by Edwin Southern. Detection of RNA is termed northern blotting and the detection of posttranslational modification of protein is termed “Eastern blotting”.

Media Preparation Before mixing medium, determine the quantity of each medium and the amount of plant growth regulator needed for each experiment. Make slightly more medium than is required. A media check off list is helpful in making medium to keep track of ingredients. Remove media stock solutions and plant growth regulator stock solutions to be used from the refrigerator. Make up fresh stock solutions if adequate quantities are not available or if solutions have precipitated. Plant growth regulators stock solutions are prepared biweekly and stored at 4º C in the dark. Cytokinins (thidiazuron and BA) are dissolved in approximately KOH prior the addition of deionized water. Gibberellins and auxins (2,4-D, IBA and NAA) are dissolved in 100% ethanol prior the addition of water. Fill the volumetric flasks upto ½ to ¾ with deionized water. Add proper a of stock solution and plant growth regulators and mark each item on the check off list. Add 30 g/l sucrose, stir until dissolved. Make up the volume of the flask with deionized water and later por into a larger beaker.

Principle Orchid seeds are produced in large numbers but are exceedingly small, hence are also called “dust seeds” may be as small as 0.01 by 0.05 mm in dimension and weigh as little as 10-30 ug. Seedling development is generally a slow process in orchids, with the seedlings passing through an initial stage, termed the protocorm stage, prior to the production of distinct vegetative organs (leaves, roots, etc). The protocorms of terrestrial orchids are subterranean and entirely heterotrophic, with all of the nutritional needs of the orchid being supplied by the mycorrhizae (the symbiotic fungus). The length of the protocorm stage is variable, however, in many cases, months, or even years, elapse before leaves and roots are produced. The combination of a small seed size and fungal symbiont requirement largely precludes the propagation via conventional seed propagation protocols. Accordingly, most of the early research on the development of orchid seed germination protocols focused on in vitro germination with the seeds and fungal symbiont being sown on the same petri plate. Successful protocols were developed for several species; however, the process was complicated by the diversity of fungal symbionts required (across orchid taxa) and the difficulty in isolating and culturing these fungi. In the 1920’s the process of orchid seed propagation was revolutionized by Lewis Knudson with the demonstration that nutrient media that could be used as a substitute for fungal infection. The isolation and culture of multiple fungal strains was no longer required, and improvements in germination frequency, germination speed, and seedling vigor were generally observed. To date, successful asymbiotic germination techniques, termed ‘flasking’, have been developed for many epiphytic orchids, with several species being saved from extinction via the application of these techniques.

Principle Micropropagation is especially useful for include propagation of high value plants that grow slowly (bulbs, ferns, orchids, palms, etc), new cultivars having limited stock and high market demand (new cultivars produced via conventional breeding or genetic engineering), cultivars that are difficult to root by conventional techniques, and endangered plant species. In addition, virus elimination is possible via culture initiation from meristem explants. Also, micropropagation provides a source of sterile tissues that can be used for other tissue culture protocols, including genetic transformation. Micropropagation is possible using multiple types of cultures; however, shoot cultures are typically employed, in which case five stages to the micropropagation process are typically recognized as outlined below. Nodal explants of rose are used for culture initiation. Three media, differing in hormone content, is employed and the explants should be evaluated for differences in axillary shoot growth and proliferation.

This protocol is designed for potato stem internodes on ZIG medium, but might work for leaf pieces as well. Plant Material Cut stem internode explants from 5-6-week-old in vitro plants (or leaf explants from 3-4-week-old in vitro plants) grown on 15 ml PROP in 25 x 150 mm test tubes. Day 1 Streak out desired Agrobacterium strain on YEP + antibiotic to obtain a single colony. Grow at 28 C for 2 days. Alternatively, skip this step and use frozen stock for next step. Day 3 Use a single Agrobacterium colony to inoculate 20 ml YEP + antibiotic in 250-ml flask. Grow overnight at 28 C with vigorous shaking (~200 rpm). Some Agrobacterium strains require an additional day to reach sufficient density.

After preparing the desired DNA molecule using E.coli, the plasmid can be transferred into Agrobacterium by this method. Efficiency is low (~103 transformants per mg of DNA) but the method is reliable and very rapid. This method also eliminates much of the plasmid rearrangement that occurs during triparental mating. Procedure Grow an Agribacterium strain containing an appropriate helper Ti plasmid in 5 ml of LB medium 16 hours at 28ºC. Add 2 ml of the culture to 50 ml LB medium in a 250 ml flask and shake (~250 RPM) at 28ºC until an OD600 of 0.5 to 1.0 is obtained. Chill the culture on ice. Centrifuge the cell suspension at 1000 g for 30 minutes at 4ºC. Discard the supernatant. Resuspend the cells in 1 ml of 20 mM CaCl2 (ice cold). Dispense 100 mL aliquots into pre-chilled 1.5 ml tubes. Add about 1 µg of the desired plasmid DNA (pure or crude) to the cells in each tube.

Appendices APPENDIX-I : Solutions & Buffers Solutions 10X ABI TBE 216 g Tris base 110 g boric acid 16.6 g EDTA Add water to 2 liters.