ANG 111 Summer EXPERIMENT 1: CLONING June 23 July 14

ANG 111 Summer 2009 EXPERIMENT 1: CLONING June 23 July 14 Lab report for this experiment is due JULY 17, 2009 BY 5:00 pm. Late write-ups will be severely penalized. NOTE: IT IS IMPORTANT TO READ THESE LABORATORY DIRECTIONS PRIOR TO COMING TO LAB. YOU ARE EXPECTED TO HAVE THOUGHT ABOUT THE MATERIAL IN THE DIRECTIONS. THERE ARE QUESTIONS INTERSPERSED IN THE DIRECTIONS. YOU WILL NOT GET THE FULL BENEFIT OF THE LAB IF YOU DO NOT READ THEM IN ADVANCE (NOR WILL YOU GET FULL CREDIT ON THE QUIZZES THAT PRECEDE EACH LAB). Purpose of the exercise: The first experiment is designed to give the technical experience necessary for cloning genes. The experiment is designed to subclone a region of DNA; that is, you will be taking a specific piece of the DNA from one vector and moving it to another, more useful, vector. Overview: Cloning and subcloning are tools used to isolate and propagate a gene or portion of a gene of interest (affectionately referred to as Your Favorite Gene or YFG). The clone can be amplified to obtain large quantities of DNA to study the function and properties of the gene in vivo or in vitro. The fragments of genes that you clone can also be used to construct new genes for either study or for use in changing the phenotype of a plant or animal. Background: The DNA fragment with YFG is ligated (covalently linked using the enzyme ligase) into a plasmid vector. The plasmid vector provides the origin of replication, which is necessary for replicating YFG in E. coli or another suitable bacterial host. Bacterial cells (typically E. coli strains) provide the essential proteins that act on the origin of replication of the plasmid to replicate the cloned DNA. Bacterial cells are transformed with the ligated plasmids, i.e. the plasmid DNA passes through the bacterial cell wall and enters the cell. In a nutrient rich culture medium (such as LB), the bacterial cells replicate exponentially and the plasmid clone replicates extrachromosomally inside the cell. A technique called a plasmid preparation, or miniprep if only small quantities of the plasmid DNA are prepared, is used to isolate the replicated plasmid DNA from within the cell. Rationale: We are subcloning a portion of a DNA construct ( construct referring to a DNA molecule created in the laboratory) that was explicitly made for the generation of transgenic goats. This construct consists of the bovine -lactoglobulin (BLG) promoter (1267 base pairs long) fused to the human lysozyme (HLZ) gene (540 bp cdna) and BLG 3 flanking sequences (3180 base pairs (bp) or 3.18 kilobases (kb) long). Currently this DNA construct is in the plasmid vector pgem-t Easy and is referred to as pblg-hlz (Figure 1). We wish to subclone the promoter and HLZ coding region (a 1.8 kb SacII-EcoRV fragment) into a plasmid vector that is more useful to us, a Bluescript plasmid vector (pbs). 1

OVERVIEW OF THE PROCEDURES IN THIS EXPERIMENT: Large-scale preparations of the pblg-hlz and Bluescript plasmids will be provided (Figure 1). Question: What size is each of the two plasmids? Where would you expect to see them on an electrophoresis gel? Cut each plasmid DNA preparation first with SacII and then with EcoRV restriction enzymes (Protocol 1). Question: How will we know if we successfully cut the plasmids? Why do we need to cut the Bluescript (the target vector)? Isolate the 1.8 kb fragment of the promoter/hlz coding region from pblg-hlz (Protocol 2). Question: Why must the fragment (or insert ) be isolated? Estimate the concentration of the cut Bluescript plasmid and the isolated HLZ fragment (Protocol 3). Ligate the HLZ insert into the Bluescript overnight at 16 o C (Protocol 4). Check the ligation product on a gel (Protocol 4). Question: Why do we need to do this? Transform E. coli (strain DH5 ) cells with the ligation product (Protocol 5). Plate the transformed bacterial cells on plates containing the antibiotic ampicillin (amp) as a selection agent and grow at 37 o C overnight (Protocol 6). Question: What do we mean by selection agent? Pick individual colonies (= clones) and grow in suspension culture overnight in medium containing ampicillin (Protocol 7). Question: Why do we need to maintain the selection if we selected in step 8? From the suspension cultures, isolate plasmids containing the HLZ coding region DNA using a miniprep procedure (Protocol 8). Perform restriction enzyme digestions and Southern blot on isolated plasmid DNA (Protocols 9 and 10). Prepare a digoxygenin labeled probe and hybridize with the Southern blot membrane (Protocol 11) to verify that the correct fragment has been cloned. Question: Why don t we just assume the clones that grew in the selective medium contain exactly what we want? 2

pgem-t Easy SacII 1.3 kb XhoI SacII NotI PstI EcoRV KpnI pblg-hlz (8 kb) 540 bp XhoI EcoRV 1.8 kb pbs (2.9 kb) ClaI SphI 1.3 kb Figure 1. Plasmids used in Experiment 1. Bovine -lactoglobulin (BLG) promoter (1267 bp) Human Lysozyme cdna (HLZ) promoter (540 bp) Bovine -lactoglobulin (BLG) coding region, exons 2 and 3 (1831 bp) Bovine -lactoglobulin (BLG) coding and 3 flanking region (1349 bp) pgem-t Easy (3018 bp) 3

Protocol 1- Digestion of plasmids with SacII and EcoRV restriction enzymes JUNE 23, 2009 PROTOCOL 1a: SacII digestion 1. Digest the plasmids pblg-hlz and Bluescript with SacII in separate tubes. Add the reagents in the listed order to the plasmid DNA in a 1.5 ml microfuge tube. Change pipet tips between reagents; do not cross contaminate the reagents. Add the reagents to the bottom of the tube, and mix by gently swirling with the pipet tip. Mixing too hard can denature your enzyme! Make sure that the enzyme reaches the DNA. Enzymes are heat labile. Keep them and the reactions on ice until you are ready to initiate the digestion in step 2. Reaction tube #1 Total Volume: pblg-hlz digestion: 10 l pblg-hlz plasmid (plasmid concentration is 1 g/ l) 12 l ddh 2 0 4 l Bovine serum albumin 4 l 10X restriction enzyme buffer for SacII 10 l (1unit / l) SacII 40 l Reaction tube #2 Bluescript (pbs) digestion: 10 l Bluescript plasmid (1 g/ l) 12 l ddh 2 0 4 l Bovine serum albumin 4 l 10X restriction enzyme buffer for SacII 10 l (1unit/ l) SacII Total Volume 40 l Question: How much (in g) pblg-hlz plasmid is in the reaction? Question: Why were 12 l of double distilled H 2 0 added in tube #1? Question: Why do we add the enzyme last? 2. Close the tubes, LABEL each tube with a description of the contents AND your section and group numbers, gently mix the contents of the tube, spin in the centrifuge for 15 seconds to bring contents to bottom of tube, and then digest for at least 1 hour in a water bath set at 37 o C. While the digestion is taking place, look at the plasmid maps (Figure 1). Questions: Where will the DNA be cut in the pblg-hlz plasmid? Where will it be cut in the Bluescript plasmid? Will you get one piece or two? Draw out what you expect the successfully cut plasmids to look like. What size band(s) do you expect to see on your gel for each plasmid? Each group needs to make a 0.8% agarose mini-gel for the next step. Use the gel comb that makes 8 wells. 4

3. Check the plasmid digests on the agarose mini-gel. a. Label 5 small microfuge tubes 1-5. b. To tube #1, add 5 l of the pblg-hlz digestion reaction, 1 l sample dye and 4 l ddh 2 0. c. To tube #2, add 5 l of the pbs digestion reaction, 1 l sample dye and 4 l ddh 2 0. d. To tube #3, add 1 l of undigested pblg-hlz, 1 l sample dye and 8 l ddh 2 0. e. To tube #4, add 0.5 l of undigested pbs, 1 l sample dye and 8.5 l ddh 2 0. f. To tube #5, add 10 l of DNA marker (1 kb) containing sample dye. Question: Why are you running uncut plasmid? Question: What do you expect to see on the gel for the uncut plasmid samples? Question: Why can you expect a difference in migration distance between linearized and circular plasmids on agarose gels? This figure represents your gel lanes: 1 2 3 4 5 6 7 8 Before loading the gel, make a list of what sample you will put into each well: 1) 2) 3) 4) 5) 6) 7) 8) g. Run the gel with its samples at 220 volts for 35 minutes (min). h. View the gel with the ultraviolet light box/ ChemiImager - save and print the image. 4. The digested DNA will be concentrated by precipitation in a solution with ethanol and salt. 5

Question: What is the volume of solution remaining in each of your digested DNA samples? Digested pblg-hlz l pbs l Precipitate each digested plasmid DNA by adding 1/10 of the sample volume of 3 M NaOAc - and 2.5 times the sample volume of cold, 100% ethanol. Volume in these cases refers to the reaction volume you are precipitating. For example, if there is 35 l of digested pblg-hlz, add 87.5 l ethanol and 3.5 l NaOAc. Allow the DNA to precipitate at -20 o C until the next lab. JUNE 24, 2009 Protocol 1 (continued) PROTOCOL 1b: EcoRV digestion 5. Spin the sample tubes containing the digested plasmid DNAs in a microfuge at 4 o C for 10 min at maximum speed. Since this is a fixed rotor centrifuge, note where you should expect to find the pellet- be consistent with how you insert the tubes. Carefully remove the ethanol supernatant without dislodging the DNA pellet. Rinse pellet by adding 100 l of cold 70% ethanol, spin 2 min in the microfuge and then remove the supernatant. Let the DNA dry by turning the tube upside down and leaving at room temperature for about 10-15 minutes. Blotting out liquid with a tissue is sometimes helpful. Resuspend the pellet in 20 l ddh 2 O. 6. Digest the samples as in step 1, but with EcoRV replacing the SacII restriction enzyme and substituting the appropriate buffer. puc18-gh9 digestion: 20 l pblg-hlz plasmid (~10 g) cut with SaII and precipitated 2 l ddh 2 0 4 l Bovine serum albumin 4 l 10X buffer for EcoRV 10 l (1 unit/ l) EcoRV Total Vol. 40 l Bluescript digestion: 20 l Bluescript plasmid (~10 g) cut with SacII and precipitated 2 l ddh 2 0 4 l bovine serum albumin 4 l 10X buffer for EcoRV 10 l (1unit/ l) EcoRV Total Vol. 40 l 7. Digest for at least 1 hour at 37 o C. While DNA is digesting, each group should prepare a mini-gel for evaluating the digestions. 6

8. Repeat step 3 from previous lab to confirm appropriate cutting by EcoRV What controls do you need? Plan the loading ahead while your DNA is digesting. Question: What band pattern do you expect if the digestions are complete? 9. Precipitate the digested pbs (but not the pblg-hlz) as described in Step 4 from your previous lab. 10. If the pblg-hlz plasmid is completely digested, place at -20 o C until next lab. If additional digestion is required, tell your instructor or TA. Remember to label each tube with its contents and your section and group numbers. JUNE 29, 2008 Protocol 2 PROTOCOL 2: Purification of insert fragment We will use a Qiagen Gel Extraction Kit to isolate the insert. In this procedure the DNA is bound to a specific matrix, washed to remove contaminants, and eluted as purified DNA. 1. Each group will pour a 0.8% agarose gel using a 6 or 8-well comb. 2. Add 1/10 th volume of loading dye to your double-digested pblg-hlz DNA. Load marker DNA into the first lane and then skip a lane between the marker and the sample lanes. Load the entire volume of the doubly digested pblg-hlz plasmid into two wells, side by side. Run the gel at 200 volts for 30 min. Question: Why are we only running the pblg-hlz sample? 3. While the gel is running, weigh a 1.5 ml microfuge tube. Place the electrophoresed gel on the UV light box. Cut a slit in the gel just ahead and behind the band of interest. Remove the gel slice with a spatula. NOTE: It is important to keep the size of the agarose piece with your sample small. Be sure to wear goggles and minimize time on the UV box! Why? Break the gel slice into pieces and put up to 400 mg of the gel into the weighed 1.5-ml tube. Add 300 l of Buffer QG for every 100 mg of gel. Buffer QG is a solution that will dissolve the gel. 4. Incubate at 50ºC for 10 min. Mix every 3 min by inverting or vortexing tube to aid gel dissolution. If there are still gel pieces in the solution, continue incubation and mixing until all of the gel pieces have completely dissolved. 5. After the gel pieces have dissolved, check the color of the mixture. If the color is yellow (similar to Buffer QG), continue to Step 6. If the color is orange or violet, add 10 l of 3M sodium acetate, ph 5.0 and mix. The color of the mixture should return to yellow. Buffer QG contains a ph indicator. The ph of the solution must be 7.5 in order to get efficient absorption of the DNA to the matrix for purification. 7

6. Place a QIAquick spin column into a 2-ml collection tube. Pipet up to 800 l of the solution containing the dissolved gel into the spin column. Centrifuge the mixture in a microcentrifuge at 14,000 rpm for 1 min. Make sure you balance your sample in the centrifuge! Discard the flow-through. Your DNA should be stuck to the matrix in the column. If your sample volume was more than 800 l, load the rest of the sample and spin the column again. 7. Place the cartridge back into the empty 2-ml collection tube. Add 500 l of Buffer QG to the column and centrifuge at 14,000 rpm for 1 min. Discard the flow-through. Your DNA should still be stuck to the column. This step removes any traces of agarose. 8. Place the spin cartridge back into the empty 2-ml collection tube. Add 750 l of Buffer PE to the spin column and incubate for 5 min at room temperature. Centrifuge at 14,000 rpm for 1 min. Discard the flow-through. Centrifuge again at 14,000 rpm for 1 min to remove residual buffer. Your DNA should still be stuck to the column. 9. Place the spin cartridge into a NEW 1.5 ml recovery tube. Add 30 l of Buffer EB directly into the center of the spin column. Incubate for 1 min at room temperature, then centrifuge at 14,000 rpm for 1 min. Your DNA will be off the cartridge and in the tube. Save the DNA! Label the tubes clearly and store at -20 0 C until the next lab. 10. Spin down the precipitated pbs from your previous lab session as described in Protocol 1, Step 5. Rinse, dry, and resuspend in 20 l dd water. Store at -20 0 C until the next lab. JUNE 30, 2009 Protocols 3 and 4 PROTOCOL 3: Estimating DNA concentration by SYBR Safe staining We will approximate the concentration of our DNA fragment by comparing it to a DNA marker of known concentration on a gel. 1. Each group should prepare a 0.8% agarose mini gel. 2. Calculate how many nanograms (ng) of your purified DNA fragment you would have if your recovery were 50% by going through the following calculations: You know how much pblg-hlz DNA was used at the beginning, therefore you should be able to estimate the amount of insert DNA that you have as outlined below. In what follows, it is important to distinguish between pblg-hlz plasmid (= 8 kb) and the promoter/hlz fragment (= 1.8 kb): You can calculate how much of the 10 µg pblg-hlz plasmid DNA that you started with was accounted for by the 1.8 kb promoter/hlz fragment as follows: 8

The pgem-t Easy vector alone is 3018 bp. The total size of the pblg-hlz plasmid is 8005 bp (1257 bp BLG promoter + 540 bp HLZ coding region + 1831 bp BLG coding region + 1349 bp BLG 3 flanking DNA + 3018 vector). The HLZ SacII-EcoRV fragment is 1884/8005 or ~24% of the weight of the pblg-hlz (assuming about the same distribution of ATGC along the entire 8005 bp). Therefore, you started with about 2.4 µg of HLZ fragment. BUT, some of these 2.4 µg were used for gels to verify that the digestions worked: 10 µg (=10 µl) pblg-hlz plasmid were used as starting material for SaII digestion. The total reaction volume for this digestion was 40 µl but 5 µl were used (lost) for the gel to verify digestion efficiency. Questions: How much (in µg) HLZ fragment was present in the remaining 35 µl after SacII digestion assuming maximum recovery after DNA precipitation? µg How much (in µg) pbs fragment was present in the remaining 35 µl after SacII digestion assuming maximum recovery after DNA precipitation? µg The total reaction volume for the EcoRV digestion was also 40 µl and again 5 µl were used (lost) for the gel to verify digestion efficiency. Questions: How much (in µg) HLZ fragment was present in the remaining 35 µl after EcoRV digestion assuming maximum recovery after DNA precipitation? µg How much (in µg) of pbs fragment was present in the remaining 35 µl after EcoRV digestion assuming maximum recovery after DNA precipitation? µg With the Qiagen band isolation technique, if we assume a recovery constant of approximately 40%, we should have about ng of HLZ DNA. If the total volume of solution containing our fragment after the purification is 30 l, what is the estimated concentration of the fragment? ng/ l What volume contains 0.2 µg (200 ng) of HLZ fragment? l What volume contains 200 ng of pbs? l 3. Mix in a microfuge tube the volume of solution with your estimated 200 ng of the HLZ fragment and 1 l of sample dye. Bring the final volume to 10 l with water. 4. In another tube, add the volume of solution with your estimated 200 ng of pbs and 1 l sample dye. Bring the final volume to 10 l with water. 5. Load both samples on the gel along with 10 l of your marker DNA. Run the gel at 200 volts for 35 minutes, then look at the gel with the UV light/ ChemiImager. 9

SYBR Safe binds to the DNA in the gel, and under UV light when bound to DNA, it fluoresces with an intensity proportional to the amount of DNA present. How much DNA is in your marker lane band that is nearest to the HLZ band? ng How much DNA is in your marker lane band that is nearest to the pbs band? ng By comparison, how much DNA do you estimate is in your HLZ band? ng How much DNA is in your pbs band? ng What is the actual concentration of HLZ in your sample? ng/ l What is the actual concentration of pbs in your sample? ng/ l Question: How do these actual values compare to the expected recoveries calculated on the previous page? Calculate the amount of HLZ fragment that you have left in your microfuge tube. ng Calculate the amount of pbs that you have left in your microfuge tube. ng PROTOCOL 4: Ligation of DNA 1. You want to ligate the purified HLZ fragment to the digested pbs. It works best to use 3x the number of insert molecules as acceptor (vector) molecules. The number of molecules depends on the size of the molecule as well as the weight in nanograms. That is, if you have a 100 bp fragment and a 300 bp fragment at equal concentrations (ng/ l), the 100 bp fragment has 3 times as many molecules and 3 times as many ends as the 300 bp fragment. pbs is 2960 bp. The HLZ fragment is approximately 66% of the length of the pbs. To have 3x the number of HLZ molecules as pbs molecules, we want the HLZ fragment to be present at 3 x 66% (= twice) the concentration of the pbs. In other words, 200 ng of HLZ insert would need to be ligated with 200 ng / 3 / 0.66 or 100 ng of the pbs. This is about l of your digested pbs. 2. T4 DNA ligase comes commercially packaged with a tube of 10X ligation buffer. The total volume of the ligation reaction will be 20 l. Therefore, you must add 2 l of 10X ligation buffer and 1 l of T4 DNA ligase. So you have 17 l left to add your two DNA fragments. In general, the combined amounts of your two DNA fragments should be at a minimum of 300 ng and maximum of 1 g in the following reaction. Add water to make up the volume if needed. Remember change your tips, and add the heat labile ligase last. Label a 500 l tube HLZ Ligation and add the following components: 10

HLZ Ligation: Final volume 2.0 l 10X ligation buffer l isolated HLZ fragment (use above estimates- at least 20 ng) l Bluescript cloning vector (use above estimates- at least 10 ng) l ddh 2 0 1 l T4 DNA ligase 20 l In another tube, prepare a second ligation mixture. This is a control for how well the double digestion really worked in cleaving pbs. Label this reaction Ligation Control Ligation Control: Final volume 2.0 l 10X ligation buffer l digested Bluescript cloning vector. (Use the same volume here as you used above) l ddh 2 0 1 l T4 DNA ligase 20 l Question: Why is this a control for how well the restriction digests worked? 3. Keep the reaction on ice until you are ready to incubate the ligation mixture at 16 o C, overnight. SAVE THE REMAINING DIGESTED pbs FOR THE NEXT LAB. JULY 1, 2009 Protocols 4, 5 and 6 PROTOCOL 4 (continued) 4. If sufficient DNA was recovered, we should be able to see the ligation reaction product on an agarose gel. Each group will make mini gels of 0.8% agarose. In the first lane load the marker; the next lanes will be for your ligation products. Take 5 l of each of the ligation reactions and add 1 l of sample dye + 4 l water. Run the gel for 25 min at 200 volts. DO NOT ADD SAMPLE DYE TO THE REST OF THE LIGATION REACTION! PROTOCOL 5: Transformation of E.coli DH5 Cells 1. Label five 15 ml white-top tubes clearly (A, B, C, D, E). Add 50 l of E. Coli DH5 competent cells to each tube. Keep cells on ice at all times. 11

A. In Tube A, add the remaining contents of the HLZ Ligation reaction to the competent cells. The other four tubes will be used for controls as follows: B. To Tube B, add 15 l ddh 2 0. C. To Tube C, add your digested but UNLIGATED pbs. Add the same quantity of pbs DNA that you used in the ligation reactions. D. To Tube D, add the remaining contents of the Ligation Control reaction. E. To Tube E, add the same amount (µg) of UNDIGESTED pbs DNA that you used in the ligation reactions. [Hint: you may find it useful to dilute the DNA] Question: What is the purpose of each of the controls? What do you expect to see on the bacterial plates for each control when plating them all on LB-Amp plates? 2. Incubate the DNA and competent cells on ice for 30 minutes. 3. Heat shock the 5 transformation tubes by incubating for 30 sec in a 42 o C water bath. NOTE- the time at heat shock temp is important! 4. Remove the tubes from the heat shock, and place the reaction on ice for 5 min. 5. Add 1 ml of LB broth to each of the 5 tubes containing the competent cells and incubate at 37 o C with shaking for 1 hour. PROTOCOL 6: Plating the transformation product using sterile technique The contents of each tube will be plated on LB (Luria Broth) or LB-Amp plates using 500 l of reaction mix per plate. The Bluescript vector has ampicillin resistance, so only cells containing this vector should grow on LB-amp plates. To help maintain a sterile environment always work near a bunsen burner flame. Watch your hair, shirtsleeves, and notebooks when working near open flames. DO NOT leave a burner going if you are not immediately adjacent to it. 1. Get seven plates: 6 LB + AMP and one LB only. As instructed below, you will add X- gal (a chromogenic compound that is cleaved by the lac-z gene product to form a blue color) to the six LB+AMP plates. The LB-only plate will be used for the transformation control using water (Tube B). Question: Why does the plate for this control use LB only, with no added Amp? Do you need to add X-gal to this plate? Clearly label each plate by writing on the top of the lid using a black marker and 12

SMALL print. Designate exactly what is on the plate, including the selective reagents, cells, DNA. Using sterile technique, first mix 240 l X-Gal solution (20 mg/ml) with 360 l LB in a sterile tube. Add 100 l of the diluted X-Gal solution to the middle of the LB+AMP agar and immediately spread over the plate covering the entire top using the L-shaped glass spreader. Flame the spreader and allow it to cool before each use. Repeat for each of the six plates. Let dry while you are getting situated. 2. Using fresh pipette tips, add the indicated amount of transformed cell solution to the appropriately labeled plates. Tube A: plate on two LB-Amp/X-Gal plates (250 l on one, 500 l on the other) Tube B: plate on one LB-Amp/X-Gal plate and one LB only plate (500 l each) Tube C: plate on one LB-Amp/X-Gal plate (500 l) Tube D: plate on one LB-Amp/X-Gal plate (500 l) Tube E: plate on one LB-Amp/X-Gal plate (500 l) Keep the plates covered as much as possible to prevent contamination. Spread the ligation reaction products on the plates with the L-shaped glass spreader until no liquid can be seen on the agar s surface. Be sure to flame the spreader between uses with the different ligation products. 3. Invert the plates and place in an incubator at 37 o C, overnight. JULY 6, 2009 Protocol 7 PROTOC0L 7: Picking individual colonies (clones) using sterile technique Count and record the number of white and blue colonies on each plate. Colonies that appear to contain the appropriate subcloned HLZ will be selected and verified. Question: How do we recognize those colonies that should have the subcloned DNA in them? What do each of the controls tell us? 1. Add 5 ml of LB broth containing ampicillin to each of 4 culture tubes using a sterile pipette and working near a flame. Question: Why do we use LB containing ampicillin? 2. Using a sterile yellow pipette tip and gloves, scrape the tip on an individual colony from the HLZ ligation plates, then drop the tip into the culture tube and cap. Label your tubes 1-4 and put group number and initials on each of them. Pick from the middle of the colony to avoid satellite colonies. 3. After you have picked your colonies put the tubes in a rack in a 37 o C shaking incubator overnight. 13

JULY 7, 2009 Protocols 8 and 9 PROTOCOL 8: DNA miniprep- Isolating plasmid DNA from E.coli by alkaline lysis with SDS 1. From each of the 4 overnight cultures, pour 1.5 2.0 ml of the cell suspension into a 2.0 ml microfuge tube. Make sure the cells are in suspension before pouring (you can gently shake the tube). Label your microfuge tubes. 2. Centrifuge at room temperature for 30 seconds at maximum speed. Pipette out the supernatant being careful to not disturb the cell pellet. Remove as much medium as you can! 3. Add 100 l of ice cold Solution I (recipe for Solutions I, II, and III are at the end of this protocol) to each microfuge tube, vortex vigorously to ensure the bacterial cell pellet is dislodged and resuspended. Incubate on ice for 5 minutes. 4. Add 200 l of solution II, and invert several times until the white pellet is completely dispersed. Do not vortex. Incubate on ice for 5 minutes. 5. Add 150 l of ice-cold Solution III, and invert several times to mix. Incubate on ice for 5 minutes. 6. Centrifuge at maximum speed at room temperature for 2 min. 7. Transfer the supernatant to a 1.5 ml microfuge tube. Add 450 l phenol-chcl 3 (yellow) solution. Be careful! Phenol can cause serious skin burns. Mix by vortexing. 8. Centrifuge at maximum speed for 2 min. The upper layer contains the soluble contents of the lysed E. coli including the plasmid DNA. This step is referred to as a protein extraction step since we are removing proteins (mostly at interface between the two layers) from our DNA solution. 9. Transfer the aqueous (top) phase, which contains the DNA, to a new 1.5 ml microfuge tube. Be careful not to transfer the interphase as it contains the denatured proteins! Add 0.9 ml of cold 100% EtOH and mix well. Question: Why do we add EtOH at this step? Why don t we add NaOAc as we have in the past? 10. To aid precipitation of the DNA, put your tubes in a dry ice-etoh bath for 20 min. This is a speedy way to precipitate DNA, but it isn t as efficient in terms of DNA recovery. 11. Centrifuge at 4 o C for 5 minutes at maximum speed. Remember to note where you expect your pellet to be found. 12. Remove the supernatant with a pipet. Discard supernatant, save pellet. 14

13. Rinse the pellet with 1.0 ml of cold 70% EtOH, then spin the pellet for 2 min. 14. Pipet off and discard the 70% EtOH. Dry the pellet by inverting and leaving at room temperature for about 10 min. 15. Add 50 l of RNaseA-TE to resuspend the pellet. Question: Why do we add RNaseA to our DNA preparation? DNA Isolation (Miniprep) Solutions (these will be made up for you): Solution I 25mM Tris-HCl ph 8 10 mm EDTA 50 mm glucose Solution II 0.2 N NaOH (freshly diluted from a 10 N stock) 1% SDS Solution III 60 ml 5 M Potassium acetate 11. 5 ml glacial acetic acid 28.5 ml dd H 2 0 PROTOCOL 9: Restriction enzyme double digestion using SacI and EcoRV. For three of your four minipreps, set up the following reaction mix: Total volume 10 l colony miniprep plasmid DNA 2 l 10X Enzyme buffer (Fermentas FastDigest ) 2 l water 1 l SacI (1U/ l) 1 µl EcoRV (1U/ µl) 20 l Put the samples in 37 C incubator for 10 minutes to digest. Store the samples at 20 C after they have completed the incubation. Save the rest of the uncut plasmid of the same minipreps to use as controls for next time! July 8, 2009 Protocols 10-1 AND 10-2 PROTOCOL 10, PART 1: Gel electrophoresis for transfer and to check digests 1. Each group should prepare a 0.8% agarose gel with at least 8 wells. 2. Add sample dye to your digested samples (2 l dye/sample). 15

3. For each sample, prepare a corresponding amount of undigested miniprep DNA by placing the same amount of sample you used in your double digest in a new tube with sample dye (10 l undigested plasmid + 10 l H 2 0 + 2 l dye). 4. You will be provided with 10 l (100 ng) of purified HLZ fragment to use as a positive control for the Southern blot. Add 1 l sample dye to the sample. 5. Load 10 µl of each sample, digested and undigested. In Lane 1, load the 1 kb marker. In lanes 2-7 load your DIGESTED and UNDIGESTED samples. In lane 8, load the purified HLZ. While the gel is running think about what bands you expect when the correct clone is cleaved. Question: Why do you want to run an uncut version of each isolated colony s DNA? 6. After running the gel for 45 min at 250V, examine it under a UV light and take a picture. Include a fluorescent ruler along side the marker lane. Question: Why do we include the ruler? PROTOCOL 10, PART 2: DNA transfer (Alkaline Transfer) Transfer the gel to a glass dish. Cut off and discard the marker lane and unused areas of the gel including the section above wells with a razor. Soak the gel for 30 min with gentle shaking in enough 0.4M NaOH to cover the gel. Prepare membrane for transfer: Cut membrane to size of gel. Also cut 6 sheets of thick blotting paper to the same size as the membrane. (NOTE: Use gloves and blunt forceps) Soak the nylon membrane in distilled water followed by immersion in 0.4M NaOH for at least 5 minutes. Set up semi-dry blot transfer (DNA will be transferred upward by capillary transfer). TA will do next steps for you: After at least 4 hours, remove membrane and mark it in pencil to show orientation, positions of lanes, and an identifier for your group. Wash in 2 x SSC, then allow to dry and crosslink using UV light. 16

July 13, 2009 Protocol 11 PROTOCOL 11: Labeling DNA probe and hybridization We will use a nonradioactively nucleotide (digoxigenin-dutp) to label our probe and a detection system provided by Roche Molecular Biochemicals (see Figure 2). Pre-hybridize membranes with 20ml of Hybridization Solution (1X DIG Easy Hyb TM ) at 68 o C for at least 1 hour, in hybridization container covered with saran wrap to minimize evaporation. Use the shaking incubator set at 68 o C. While the membranes pre-hybridize, label your probe (purified ogh fragment) with DIGdUTP. We will prepare our labeled probe by random-primed labeling. We will detect hybridization of our probe to the samples on the membrane by chemiluminescence using CDP-Star substrate (Figure 2) and the ChemiImager. 1. Add between 100 ng and 1 g of your template DNA (purified HLZ fragment) to a 1.5 ml microfuge tube and bring the total volume to 16 l by adding sterile doubledistilled water. Calculate how much volume you need based on the concentration of your fragment. 2. Denature DNA in a boiling water bath for 10 min, then quick-chill on ice for 1 min. 3. Add 4 µl of pre-mixed 1X DIG-High Prime solution (containing random hexanucleotide primers, dntps (incl. DIG-dUTP), and Klenow enzyme) to the 16 l of denatured template. 4. Mix by tapping the tube, centrifuge very briefly, then incubate 1 h at 37 C. 5. Stop the reaction by adding 2 µl of 0.2M EDTA (ph8.0). 6. Precipitate probe by adding 2.5 µl 4M LiCl and 75 µl 100% ETOH. Leave 30 min at - 70 o C. Question: Why is it important to precipitate your labeled probe? 7. Centrifuge 10 min, pipette off supernatant, wash pellet with 70% ETOH (50-100 µl) and dry. 8. Resuspend in 50 µl TE. 9. Heat the DIG-labeled probe in a boiling water bath for 5 min, quick-chill on ice for 1 min then centrifuge for 10 sec to collect the contents to the bottom of the tube. Question: Why do you need to denature the probe? 10. Leave the pre-hybridization solution on the membrane and add 5ml of fresh 1x DIG 17

Easy Hyb buffer to the membrane. Tilt the tray to one side and add all (50 l) of the denatured probe to the solution. Mix well to evenly distribute the DIG-labeled probe in the resulting 25 ml solution. 11. Cover the container well to minimize evaporation and hybridize at 68 o C with shaking for at least 6 hours. The next steps will be done for you (This procedure washes off excess probe that may be on the membrane): Wash membranes twice for 5 min at room temp with 50 ml 2 x SSC, 0.1% SDS. Wash membranes twice for 15 min at 68 o C with 50 ml 0.1 x SSC, 0.1% SDS. Question: Why is it necessary to wash the excess probe off? Store membranes in 2X SSC until next lab at 4 C. JULY 14, 2009 Protocol 12 PROTOCOL 12: Immunodetection of Southern Blot All incubations are done at room temperature. The volumes are calculated for a membrane size of 5 x 10 cm = 50 cm 2 : Wash membranes in 50 ml of 1X Washing Buffer (Buffer 1) with gentle mixing for 1 min. Pour off Buffer 1 and block membranes for 30 min by incubation in 50 ml of 1X Blocking Buffer (Buffer 2) with shaking. Dilute antidigoxigenin alkaline phosphatase (AP) conjugate 1:5000 in 1X Blocking Buffer (Buffer 2) immediately before use. Pour off blocking buffer and incubate membranes in 20 ml of the diluted mixture for 30 min with shaking. Wash membrane twice for 15 min each time with 50ml of 1X Washing Buffer (Buffer 1) with shaking. This should remove unbound antibody conjugate. Question: Why is it necessary to remove excess antibody conjugate? Wash membranes for 2 min with 20 ml 1X Detection Buffer (Buffer 3) with shaking. Place membrane with DNA side facing up on saran-wrap and apply 1 ml CSPD ready-touse. Immediately fold the saran-wrap over the membrane to spread the substrate evenly and without air bubbles over the membrane. Incubate for 5 min. Squeeze out excess liquid and seal the edges of the saran-wrap sandwich with tape. Important: Avoid drying of the membrane during exposure since this will result in dark background. Incubate the damp membrane for 10 min in a 37 C incubator. 18

Expose to X-ray film initially for 5-20 min. Luminescence continues for at least 48 hours. The signal increases in the first few hours after initiation of the detection reaction until it reaches a plateau where signal intensity remains almost constant during the next 24-48 hours. Multiple exposures can be taken to achieve the desired signal strength. Buffers Buffer 1: Wash Buffer 0.1M Maleic Acid 0.15 M NaCl 0.3% Tween-20 Buffer 2: Blocking Solution Blocking agent (non-fat dried milk) in 1X Maleic acid buffer Buffer 3: Detection Buffer 0.1 M Tris 0.1M NaCl Alkali-labile digoxigenin (DIG)-dUTP 19