IDENTIFICATON OF A TRANSFORMING PLASMID

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1 IDENTIFICATON OF A TRANSFORMING PLASMID Introduction The field of molecular genetics has resulted in a number of practical applications that have been of tremendous benefit to us. One such benefit is the ability to produce large quantities of biological materials that were previously difficult to obtain. These new production methods involve isolating the gene for the needed product and placing that gene into a rapidly reproducing organism such as a bacterium, which will then manufacture large amounts of the desired substance in a relatively short period of time. This process of altering the production capabilities of living organisms by the introduction of new genes into those organisms is commonly referred to as genetic engineering. The most common way of introducing these genes and producing these altered organisms is through the use of plasmids. Plasmids have a number of important and useful features. They are small circular DNA molecules that are naturally occurring in many bacterial species. Plasmids exist independently of the large chromosome and contain genes that encode products that are useful to the bacteria. For example, many plasmids carry one or more genes that confer resistance to a specific antibiotic. A bacterium carrying such a plasmid is able to live and multiply in the presence of that antibiotic, while cells lacking the plasmid are unable to divide. Another important feature of plasmids is a replication origin. This allows a plasmid to replicate autonomously within the bacterial cell and be present in 10s to 100s of copies. Because bacteria reproduce asexually (a process called binary fission), plasmids are passed to daughter cells. Plasmids released into the environment when a bacterial cell dies may be taken up by another bacterial cell. This type of genetic transfer is referred to as transformation. Plasmids are useful tools for the molecular biologist because they can serve as gene-carrier molecules, or vectors. A gene of interest is joined to the plasmid to form a hybrid or recombinant plasmid that is able to replicate inside bacteria. In order to prepare recombinant plasmids, it is necessary to cut the plasmid and DNA containing the gene of interest at precise locations and then to join (or ligate) the plasmid and the gene together. The cutting is accomplished by the use of enzymes called restriction endonucleases or enzymes. Hundreds of different restriction enzymes have been identified. Each one recognizes a specific sequence of 4-8 nucleotides (a restriction site) in a DNA molecule and cut at specific locations within that site. The restriction site must be present in both the plasmid and the gene to be inserted. The 73

2 restriction site of one sucli enzyme (EcoRI) is given below: cut...gaattc... >...G AATTC......CTTAAG......CTTAA G... t cut Since any given enzyme recognizes and cuts at a unique sequence, the number of cuts in any given DNA molecule is limited. Typically, the restriction sites for a given enzyme are hundreds to thousands of base pairs apart. The staggered cut of the enzyme illustrated above generates sticky, cohesive, single-strand ends. These are important in recombinant DNA work because they enable any two DNA fragments to be linked together by complementary base pairing at their ends, provided they were generated with the same restriction enzyme.. One basic procedure for generating recombinant DNA molecules is illustrated in Figure 1 on the next page. To create a recombinant plasmid, the plasmid is first cut with a restriction enzyme that has one restriction site on the plasmid. DNA containing the gene of interest is also cut with the same enzyme. The two digests are mixed; fragments cut by the enzyme can reanneal by complementary base pairing and the newly formed joints are sealed by the action of another enzyme called DNA ligase. This enzyme normally performs this fragment sealing operation during DNA replication. Direct uptake of DNA, such as a plasmid, by a bacterial cell can occur naturally when the cell is 'competent' (a state in which the cells are able to incorporate exogenous DNA). In the laboratory, competence can be artificially achieved, or induced. One such procedure involves suspending bacteria in an ice cold calcium chloride solution before adding DNA. When DNA is added to the suspension, it is given time to adsorb to the cell. The suspension is subsequently given a brief 'heat shock', then returned to chilling conditions, facilitating the transport of DNA across the membrane into the cell. Uptake of plasmid DNA in this manner is an example of transformation, and the cells are now described as transformed. For this exercise you will be presented with a broth culture of E. coli. The culture (referred to/labeled as 'A' or 'B') consists of a population of E. coli cells that have been transformed with a plasmid. Your task is to identify the transforming plasmid for your assigned culture. The transforming plasmid will be one of the following commercially available plasmids: pbad-gfpuv pblu pqe-9 puc18 74

3 Figure 1. Creating a Recombinant Plasmid GENOMIC DNA You will need information about each plasmid to help you complete your task. One type of information that you will use involves the genes that are present on the plasmid. These genes confer new phenotypes to E. coli cells that have been transformed. Identification of these new selectable phenotypes can aid in the identification of the transforming plasmid. Other useful information includes the size of the plasmid (in base pairs, or bp) and the presence and location of restriction sites. On the following page are figures of each of the four possible transforming plasmids indicating the genes that can be used as selectable marker genes. 75

4 Am 3438 bp Amp" Notes: The bla gene codes for beta-lactamase. This protein will allow bacteria to grow in the presence of annpicillin (i.e. the cells will be resistant to ampicillin). Thus the bla gene is also referred as the Amp"^ gene. Each of the four plasmids contain other important sequences, such as an origin of replication, a MCS (multiple cloning site - an engineered sequence that contains a recognition sequence for several different restriction enzymes used to insert a DNA sequence into the plasmid), regulatory sequences (e.g. promoters, etc.), among others. Only the gene(s) that are used as selectable markers are shown in the above figure. As you closely examine the selectable marker gene(s) present on these plasmids you will note that each one contains a gene that will allow transformed cells to grow in the presence of the antibiotic ampicillin. Ampicillin has been added to the growth medium to select for transformed cells. Because all four plasmids contain this selectable marker, it will not be useful in identifying the transforming plasmid of cultures A&B. 76

5 Also note that pblu and puc18 have a second selectable marker gene, the Lac Z gene. LacZ codes for an enzyme called beta-galactosidase. This enzyme catalyzes the reaction that breaks down the disaccharide lactose into its monosaccharide components (glucose and galactose). It will also catalyze the breakdown of an artificial disaccharide called X-gal to produce 'substance X' and galactose. Because substance X is blue in color, the bacterial colonies will be blue. Thus, cultures A&B can be easily screened for this phenotype by simply plating an aliquot of each culture solidified nutrient medium containing X-gal. If the resulting colonies are blue, one would predict that the cells have been transformed with pblu or puc18. If the resulting colonies are white, one would predict that one of the other two plasmids is the transforming plasmid. Since this is a simple test to perform, you will begin here. The screen for blue and white colonies will help you to narrow down the identity of the transforming plasmid. Your prediction will be tested by performing digestions of the isolated plasmid with carefully selected restriction enzymes and examining the products by agarose gel electrophoresis. During the first laboratory period you will isolate the plasmid from the remaining E. coli suspension cultures. The nucleotide sequence of each plasmid will be provided so that you can examine each for restriction sites that could be useful in their identification. Part I - Screen E. coli Cultures A and B for the Ability to Metabolize X-Gal > Wear gloves and safety glasses throughout this procedure. Do NOT leave lab with gloves on. > A/ever place a used inoculation loop on the lab bench for it will introduce unwanted bacteria into the lab environment. Place in biohazard bag. > Be sure to wash your hands before leaving lab. 1. Obtain 1 plate of nutrient agar containing ampicillin and X-gal. It is necessary to include ampicillin to maintain selection for transformed cells. Label the bottom of each plate with today's date, lab section information (instructor last name and section), your group ID, and organism ID (E. coli'^'^^^'' - A or B). 2. Obtain 1 individually wrapped plastic sterile loop. When you are ready to proceed, carefully remove one loop from its package and dip the loop into the suspension culture. Before you take the loop out of the culture, tap the loop on the inside wall of the container to break the drop out of the loop. Use only the amount of liquid that sticks to the loop to inoculate the streak plate. When inoculating a petri dish, do not lay the lid on the lab bench or touch the inside of the lid or the dish. You should open the dish just enough to perform the inoculation. 77

6 3. Streak the inoculum across about one-fourth of the plate using a zig-zag pattern, as shown in the figure on the following page (see Step 1 in the figure). Using the same sterile loop, streak into the first area several times (see Step 2 in the figure). Using the same sterile loop, streak into the second area several times (see Step 3). Invert the plate and 37 C for hours. Petri dishes are incubated in an inverted, or "upside-down," position so that any condensation that may develop will remain in the lid of the plate and will not drop onto the surface of the agar and ruin the colonies. After hours, your instructor will place the plates in the refrigerator until the next lab period. 5. {next lab period) Examine plates for bacterial growth. Growth will appear as small circular spots, or colonies. Record your observations below (e.g. note color of bacterial growth). Dispose of plates in a biohazard bag. E. CO//Culture (A or B) Colony Color on medium containing X-Gal Predicted ID of Transforming Plasmid 78

7 Part II - Isolation of Plasmid from E. coli Cultures A and B > Wear gloves and safety glasses throughout the procedure. Do NOT leave lab with gloves on. > Plasticware should be disposed of in a biohazard bag. > Read and familiarize yourself with each procedure before starting. You must follow the procedures CLOSELY. > Be sure to wash your hands before leaving lab. All steps should be carried out at room temperature. 1. Using a sterile transfer pipet, transfer approximately 2 ml of the broth culture into a sterile 2 ml microtube. The goal is to recover as many cells as possible. If the cells have been allowed to settle by gravity, do not resuspend them. Rather, remove this material first and fill the microtube with as much of the culture as possible. Be careful to avoid introducing bubbles in the microtube, for air bubbles take up volume. 2. Place the microtube in a microcentrifuge and spin at 10,000 rpm for 3 minutes. ALWAYS be sure to optically balance your tube with one from another lab group or one containing water. Also, wait for at least two other lab groups to place their tubes in the microfuge before beginning the run. 3. When the spin in completed, you should be able to see a small bacterial pellet near the bottom of the microtube. Do not proceed if you do not see a pellet. 4. Pour the supernatant into a liquid waste container at your bench. Using a micropipettor, add 250 [i\ of Buffer P1 to the pellet and resuspend the pellet. No cell clumps should be visible after resuspension of the pellet. The bacteria can be resuspended completely by vortexing or pipetting up and down until no cell clumps remain. Note: Buffer PI contains EDTA to chelate divalent cations that are cofactors of DNases; it also contains RNaseA (which will degrade RNA when the cells are lysed). 5. Add 250 yl of Buffer P2 and immediately mix the solution by inverting the tube 4-6 times. MIX GENTLY, only by inverting the tube. DO NOT VORTEX, as this will result in shearing of genomic DNA which will then contaminate your plasmid prep. NOTE: Buffer PI contains a reagent that will turn blue after adequate mixing with Buffer P2. Mixing/inverting tube 4-6 times should result in a uniformly blue suspension. Move on to step 6 even if you do not see the development of blue color after mixing/inverting tube 4-6 times. Note: Buffer P2 contains NaOH and the detergent SDS; this buffer will cause cell lysis. 6. Add 350 j of Buffer N3 and immediately and thoroughly mix by inverting the tube 4-6 times. Adequate mixing will cause the solution to turn colorless. Note: Buffer N3 neutralizes the lysate and adjusts it to high-salt binding conditions. 7. Centrifuge for 10 minutes at 13,000 rpm in a microcentrifuge. Remember, to be sure to optically balance your tube with one from another lab group. Also, wait for at least two other lab groups to place their tubes in the microfuge before beginning the run. Note: the high-salt concentration causes denatured proteins, chromosomal 79

8 DNA, cell debris, and SDS to precipitate; the smaller plasmid DNA remains in solution. 8. Carefully transfer the supernatant to a QIAprep spin column using a 100 [i\ micropipet. Be as quantitative as possible in recovery of the supernatant, BUT avoid transferring any of the white precipitate to the column. The use of a 100 jl micropipette will cause less disruption of the white precipitate than a transfer pipet or the 1000 jl micropipet. Note that the spin column is inserted/resting in a small collection tube and it does not have a 'lid'. Plasmid DNA will bind to the silica membrane at the base of the column. 9. Centrifuge collection tube with spin column at 13,000 rpm for 1 minute. Discard the flow-through (the liquid that has moved through the column and has collected in the collection tube). Replace the spin column in the collection tube. 10. Wash the spin column by adding 500 pi of Buffer PB and centrifuging at 13,000 rpm for 1 minute. Discard the flow-through and replace the spin column in the collection tube. Note: Buffer PB removes unwanted endonucleases that may be associated with the sample. 11. Wash the spin column by adding 750 pi of Buffer PE and centrifuging at 13,000 rpm for 1 minute. Discard the flow-through. Replace the spin column in the collection tube and centrifuge for an additional 1 minute to remove any residual buffer. Note: Buffer PE removes residual salts from the spin column. 12. Remove the spin column from the collection tube and seat it in a new sterile 1.5 ml microtube. Add 50 jj' of Buffer EB to the center of the silica membrane. Let the column stand for 1 minute. Do not attempt to close the microtube lid into the spin column; it will not be possible to do so; the lid should remain open. Centrifuge at 13,000 rpm for 1 minute. Note: Buffer EB is elution buffer (10 mm Tris-HCI, ph 8.5). 13. Remove the spin column from the microtube. Close the tube and label the tube with your group ID and tube contents (i.e. tentative plasmid ID). Give the tube to your instructor to store in the freezer. Part III - Determine DNA Concentration of Plasmid Samples > Wear gloves and safety glasses throughout the procedure. Do NOT leave lab with gloves on. 1. You must know the DNA concentration of your plasmid sample in jg/pl in order to set-up your restriction digestions. You will use an instrument called the NanoDrop Lite to determine the DNA concentration of your plasmid preparation. The materials you will need will be located at the instrument. 80

9 2. Your instructor will demonstrate liow to use tiiis instrument. The steps to use the instrument are given below. a. Blanking the Instrument (Note: This should be done before the first sample is measured. It is not necessary to blank between each sample, however a new blank should be taken every 30 minutes as needed) From the Home screen select DNA. Establish a blank by pipetting 1-2 j of EB buffer (from the plasmid miniprep kit) onto the bottom pedestal, lower arm and press Blank. When measurement is complete, raise the arm and wipe buffer from both the upper and lower pedestals using a dry KimiWipe. b. Measurement of Sample Pipette 1-2 JI of sample onto the bottom pedestal and press Measure. The absorbance of the sample at 260 and 280 will be taken. The absorbance at 260 is used to calculate DNA concentration (A260 is multiplied by 50 jg/ml to give concentration). Note (jg/ml = ng/pl. Be sure to record the DNA concentration for your samples. DNA concentration of your sample: Also note the A260/A280 ratio. The ratio of absorbance at 260 nm and 280 nm is used to assess the purity of DNA. A ratio of ~1.8 is generally accepted as "pure" for DNA. A significantly lower ratio may indicate protein contamination. A ratio of ~2 may indicate RNA contamination. Note this ratio for your samples also. A260/280 ratio of your sample: When measurement is complete, raise the arm and wipe the sample from both the upper and lower pedestals using a dry KimiWipe. c. Between Measurements: Wipe the sample from both the upper and lower pedestals with a clean dry, lint-free lab wipe to prevent sample carryover and avoid residue buildup. d. Last User: A final cleaning of upper and lower pedestals with dhao is recommended after the last measurement is collected as follows: Pipette 3pl of deionized water onto the bottom pedestal. DONOT USE A SQUIRT BOTTLE TO APPLY. Lower the arm and allow it to sit for approximately 2-3 minutes. Wipe away the water from both the upper and lower pedestals with a dry KimiWipe. 81

10 Part IV - Plan and Perform Digestions of Plasmid Samples with Restriction Enzymes (This can be completed outside of lab) You must determine tlie restriction digestions that will allow you to estimate the size of your plasmid and confirm its identity. A plasmid in its circular form will have a different mobility than a linear molecule of the same size (i.e. same number of bp) during gel electrophoresis. Open circular forms have a slower mobility, while supercoiled circular DNA has a faster mobility. Thus, to get the best estimate of plasmid size you will want to find an enzyme that cuts the plasmid in one location to linearize the molecule. The base sequence of each of the four plasmids is posted at your course web site. You will analyze a plasmid sequence for potential digestion products using New England BioLabs NEB Cutter program that is available at The restriction enzymes that are available for use are: Dpnl EcoRI Haelll Pst\ Ssp\ 1. To use NEB Cutter, simply copy and paste a plasmid sequence into the 'sequence box'. Indicate that the sequence is circular by clicking the button next to <circular>. Then click <Submit>. 2. A map of the circular plasmid will appear with many restriction sites indicated. To examine the digestion product(s) for a specific enzyme (or pair of enzymes), select <Custom Digest> under 'Main Options'. A list of restriction enzymes that cut your plasmid sequence will appear. Note that the recognition sequence for each enzyme is given, as well as the number or recognition sites for that enzyme in the plasmid sequence. 3. To see the outcome of a specific digestion, click the box next to the enzyme of interest (e.g. EcoRI), scroll to the bottom of the page and click <Digest>. The result of a digestion will be shown as a picture, with the restriction site(s) for the enzyme indicated on the circular plasmid. The figure on the following page shows the restriction site for HypEI (a hypothetical restriction enzyme) on the puc18 plasmid. 82

11 HypEI You can see the list of fragments generated by the specified digest (select <Fragments> under 'List') and view the results of electrophoresis (select <View gel> under 'Main Options'). To perform another digestion, click on <New custom digest> under 'Main Options'. 4. Keep the following in mind: Plan 2-3 different digestions for your plasmid. One of the digests should be a single digest (i.e. using one enzyme in the digestion reaction) that cuts the plasmid in one location. You have two possible plasmid sequences that could represent the actual transforming plasmid. Look for an enzyme that will cut each plasmid sequence in one location. This one digestion reaction will linearize your plasmid, and allow you to obtain an estimate the plasmid size. The other digest(s) can be a single or double digest (a double digest uses two different restriction enzymes in the same reaction) that will produce 2-3 fragments. Collectively, a characteristic set of fragments of known size will be generated for each of the two possible plasmid sequences. This information can be used to confirm a plasmid identity. Also keep in mind that the different fragments generated in a digestion should differ from one another by 200 bp or more in order to easily distinguished from one another by agarose gel electrophoresis. 4. Have your instructor confirm the digests you plan to perform. Be able to tell them which enzyme(s) you want to use and the size of each of the products you will generate for both possible plasmid sequences. Record this information in the table below: Enzyme(s) Fragments (in bp) generated for plasmid: Fragments (in bp) generated for plasmid: 83

12 5. If your plasmid DNA concentration is below 30 ng/pl (0.030 jg/ jl), see your instructor. They will either instruct you to work with another group that has the same plasmid to cover all of the planned digestions or provide you will additional plasmid DNA for use. You will divide your plasmid DNA sample evenly between your planned digests. Examples: If you plan 2 digests, you will have approximately 23 jl available for each digest; if you plan 3 digests, you will have approximately 15 microliters for each digest. 6. Use the formula below to determine the amount of DNA you will have in each digest. Remember 1 ng = pg. Solve for X. You want to have at least 0.45 jjg of DNA in a digest. DNA concentration (in Mg/ jl) x V (volume in pi) = X pg DNA 7. {When ready to perform the digestions) Obtain a sterile 1.5 ml microtube for each digest you have planned from the container of sterile tubes available for your lab section. Label each tube with group ID, plasmid sample (A or B) and restriction enzyme(s) to be used before proceeding. 8. Your instructor will remove the solutions you need for your digests from the freezer and place them at a 'Restriction Digestion Station' that has been set-up in the lab. Once thawed, the solutions are kept on ice during use. You will prepare your digests at this station. This is a 'community station'; DO NOT REMOVE ANY ITEMS. The only items you need to take to this station are your labeled microtubes and your plasmid DNA sample. 9. The total volume of the digestion reaction should be 50 pi. Prepare each digest as follows, adding the components in the order listed: (use a new micropipette tip to deliver each component) Add X pi nuclease free water to the microtube (take the 50 pi total volume and subtract 5 pi for the buffer, V pi for the DNA sample, and Ipl for the enzyme; this will give you the volume of water you want to use) Add 5 pi of 10X Reaction Buffer Add V pi of your plasmid DNA (enough to provide at least 0.25 pg of DNA) Add 1 ul of the restriction enzyme (note: the restriction enzyme should be added last) (50 pi total final volume) Mix components by gently flicking the tube with your finger. Follow with a 3 second spin in a mini-microfuge to bring the liquid to the bottom of the tube. 10. Incubate the restriction digests in a heating block or incubator at 37 C for 1-2 hr. 11. When the digestion incubation is completed, place your digests on ice if you are going to proceed to Part V. Place your digests in the freezer if you will complete Part V next week. Do not allow the digests to sit at room temperature. 84

13 Part V - Analyze Plasmid Digests Using Agarose Gel Electrophoresis Large organic molecules can be separated by a variety of techniques. Chromatography makes use of either paper or specially coated glass plates to separate proteins, amino acids, carbohydrates or other molecules with the aid of organic solvents. Electrophoresis is a technique that relies on differences in size and/or overall electrical charge to separate molecules in an electric field. It is most commonly used to separate either proteins or nucleic acids (both DNA and RNA). Support media most often used include cellulose acetate, starch agarose, and polyacrylamide. A liquid solution composed of one of these chemicals, when properly treated, will solidify to form a semi-solid support matrix in the shape of a slab known as a gel. Agarose gel electrophoresis is the most commonly used method for analyzing DNA fragments. The gels are porous and allow DNA fragments to move through them when a current is applied due to the natural negative charge (generated by phosphate groups) of DNA. The mobility of DNA molecules in an electric field is determined by their size and conformation (e.g. ds vs. ss, circular vs. linear, supercoiled vs. 'open', etc.). Larger fragments have greater frictional drag and are less efficient at migrating through the porous gel, thus smaller fragments have a faster mobility. The concentration of agarose used for the gel depends primarily on the size of the DNA fragments to be analyzed. Lower agarose concentrations are used to separate large DNA fragments, while high agarose concentrations allow resolution of small DNA fragments. A 1.2 % agarose gel will resolve fragments that range between 400 and 7000 basepairs. The samples to be analyzed are the digests of plasmid samples A and B. Electrophoresis of the digests will allow you to confirm your tentative plasmid ID. By including in the electrophoretic run a set of molecular weight standards (of known size), it should be possible to determine the size of any fragment detected. Preparation of an Agarose Gel > Wear gloves and safety glasses throughout the procedure. Do not leave lab with gloves on. > Read and familiarize yourself with each procedure before starting. You must follow the procedures CLOSELY. > Be sure to wash your hands before leaving lab. 1. You and your lab partner will share a gel with another lab group. Place your gel tray into a casting tray. Orient the gel mold such that the open ends of the tray are facing the rubber gaskets that line the two sides of the casting stand. Position the gel tray in the center of the casting stand. Turn the knob 85

14 at the top of the casting tray to seal the ends of the tray with the rubber gaskets. 2. Place the leveling disc in the gel tray and use the three plastic screws of the casting stand to level the apparatus. 3. Place one comb in the tray at one end. There will be groves on the gel tray for the comb to fit into. 4. Prepare 50 ml of a 1.2 % agarose solution as follows: Weigh 0.6 g of agarose into a weigh boat and transfer this to a 125 ml Erienmeyer flask. If the available balance has a maximum weighing capacity that will allow you to do so, you can weigh the agarose directly into the flask. Use a 50 ml graduated cylinder to measure 50 ml of electrophoresis buffer (Tris-acetate-EDTA, ph 8.0) and add this volume to the flask containing the agarose. Swirl contents briefly to mix. Note: The agarose will not dissolve. 5. Gently stuff a clean KimiWipe into the mouth of the flask. You will dissolve the agarose by heating it in a microwave. Begin by heating for 30 seconds. Remove and swirl flask contents - note that the agarose has not completely dissolved. Heat the solution again for 15 seconds. Watch flask carefully and stop the microwave when solution begins to boil violently and upward out of the flask. Remove from microwave and carefully observe solution - no solid particles should be visible. If necessary, heat again for 15 seconds. NOTE: If two flasks are heated at the same time, a minimum of three 30 second heating periods may be needed to melt the agarose. 6. If the volume of liquid reduces considerably during heating due to evaporation, make up to the original volume with distilled water. This will ensure that the agarose concentration is correct and that the gel and the electrophoresis buffer have the same buffer composition. 7. Allow gel solutions to cool for 2-5 minutes. (NOTE: To determine if the melted solution is cool enough to pour, touch the bottom of the flask to inside of your wrist - when it is hot (just uncomfortable, but not yet not "warm") to touch the solution is ready to pour; Do not pour an agarose solution that has begun to solidify into a gel mold). 8. Pour the liquid agarose solution into the gel mold. 9. Allow the solution to cool and solidify until it is firm (about 30 minutes). 10. Remove mold with gel from the casting tray. Remove the comb, rinse and leave it at your bench. DO NOT TRHOW THE COMB AWAY. If you will not be using your gel today, place the mold with gel into a gel storage box 86

15 containing a small amount of tank buffer. Use labeling tape to label the box with your group IDs, and store the gel at 4 C until ready to use. Analysis of Plasmid Digests Using Agarose Gel Electrophoresis > Wear gloves and safety glasses throughout the procedure. Do not leave lab with gloves on. > Read and familiarize yourself with each procedure before starting. You must follow the procedures CLOSELY. > Be sure to wash your hands before leaving lab. 1. Place the gel tray (containing an agarose gel) into the gel tank so that the wells in the gel are oriented at the cathode end of the gel tank (the end with the black electrical connection). 2. Fill the buffer reservoir with TAE buffer, covering the gel to a depth of about 2 mm (i.e. the gel is just covered with buffer). If the comb is still in the gel, remove it carefully by grasping at both sides and pulling straight up. 3. Retrieve you samples from the freezer or 37 C incubator and place them on ice. If your samples are frozen, allow them to thaw briefly at room temperature before placing them on ice. 4. To each digestion sample add 10 pi of the 5X loading dye (also called sample buffer). Use a new pipet tip for each delivery. Discard used tips into a waste beaker at this station. 5. Gently mix tube contents by flicking the tube with your finger. Spin tube(s) for a few seconds in a mini-microfuge if necessary (to bring liquid to bottom of the tube). Hold all samples on ice. 6. You should now have 55 pi of each digest ready to load. In 10 pi volumes, use a 1-10 pi micropipette to load pi of each prepared digest into a separate well. Use a new tip to load each digestion. Dispose of tips in a biohazard bag or waste collection beaker at your bench. Make a note as to the well that each sample digest was loaded into. 7. Load 10 pi of a DNA marker mix into a separate well (use a new pipet tip). The DNA marker mix contains 10 different DNA fragments ranging in size from 10,000 bp to 500 bp. Make a note as to the well that the DNA marker mix was loaded into. Return any unused DNA standard mix to your instructor. 8. Place the tank top on the gel tank and press it down in order to connect electrode wires to electrodes. NOTE: Wells of the gel should be oriented 87

16 towards the black (negative) electrode; Match the black electrode wire with the black electrode) 9. Plug the gel rig electrode wires into the power unit (match electrode wire colors with receptacle color - red with red and black with black). Turn power unit on. Select constant voltage and set display to 100 volts. Press the button with the runner symbol. If the rig was set up properly, as soon as you push the "runner" button you should begin to see small bubbles arise from the thin silver wire attached to the electrodes (the wire is covered with electrode buffer and runs across the width of the gel rig at each end). 10. Perform the electrophoresis at 100 volts until the tracking dye has migrated to 2/3 the length of the gel (approximately minutes). 11. Wearing gloves, remove the gel from the electrophoresis unit and slip the gel into a solution of ethidium bromide. There will be 3-4 gel boxes containing this solution available for lab use; each one can hold two gels at one time. Note: Ethidium bromide is a mutagen. Handel solution and items that come in contact with it with care. 12. Allow the gel to soak in the ethidium bromide soltuion for 5-10 minutes. 13. Using the spatula provided, carefully transfer the gel to a container of deionized water, and allow the gel to soak in water for 5 minutes. This will remove much of the unbound ethidium bromide from the gel, thereby reducing background fluorescence. 14. Place the gel on an ultraviolet transilluminator for examination. BE SURE TO WEAR PROTECTIVE EYEWARE WHEN USING UV LIGHT. Ethidium bromide attaches to DNA and causes it to fluoresce under UV radiation. DNA should be visible as distinct fluorescent bands. Each band represents a different DNA molecule. 15. Note the number of bands and size of each detected in each lane. If possible, take a photo of your gel with your phone or a gel documentation system. You can use the photo to collect your data. Use the fragments in the DNA ladder to estimate the size of each band in each of your digests. Note: The DNA ladder contains 10 fragments that range in size from 10 kb (kilobases) to 0.5 kb (refer to the figure on the following page). Note that 1.0 kb is equivalent to 1,000 bp (base pairs). 88

17 Kilobases -lao Figure: 10 jj of DNA ladder run on a 0.8 % agarose gel. Note that 10 fragments are present. The 3 kb (3000 bp) fragment has increased fluorescence to serve as a reference point for other fragments in the ladder. 16. Clean your bench area as follows: If your gel was stained with ethidium bromide it must be disposed of in the ethidium bromide waste container. Gels stained with gel red must be disposed of in the gel red waste container. Pour the electrophoresis buffer in your gel rig into the sink and rinse the rig with tap water. Allow the rig to dry at your bench. Discard your plasmid digests in a biohazard bag. Discard gloves in the regular trash. 17. Do the results of your digestions confirm you tentative plasmid ID? Explain why/why not. 18. Write a paragraph presenting evidence that you will use in the identification of each plasmid. 89

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