Southern Blotting MSU Potato Lab

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1 Southern Blotting MSU Potato Lab A. DNA AGAROSE GEL 1. Prepare a 1% agarose gel and load 5 μl of BMB Molecular Weight Marker DIG labeled in one lane. 2. Mix 20μg Plant Genome DNA (which has been digested with an enzyme or enzymes that will produce a fragment of DNA, encoding the desired gene, of a size between 1 and 8 kb, if possible but larger or smaller fragments should still work) with Running Dye (1X final). Centrifuge briefly and load on the gel. Be sure that a negative control (non-transgenic plant DNA digested the same) is included on the gel. As a positive control a small amount (~10ng) of the plasmid can be digested to cut out the gene (used for making the probe) and run on the gel. Since this will produce a bright signal, run this sample in the last lane; preferably separate it from a plant lane with a blank lane. With large gels, 15 cm, it is best to run overnight at about volts to ensure sharp band separation. 3. When finished, photograph the gel with a fluorescent ruler. Mark the upper left hand corner of the gel by cutting off the corner with a spatula. 4. Place the gel into a plastic tray and DEPURINATE by incubating 10 min with 0.25N HCl (250 ml or enough to cover gel completely). Agitate gently. AVOID LONG INCUBATIONS. This process cleaves DNA that allows large fragments to transfer to the nylon membrane. 5. Wash briefly twice with about 250 ml ddh DENATURE with 250 ml of SOLUTION D, 15 min. Agitate gently. 7. Repeat. 8. Wash briefly twice with 250 ml ddh 2 O. 9. NEUTRALIZE with 250 ml SOLUTION N, 15 min. Agitate gently. 10. Repeat. The gel can be left in this solution for a short time until ready to be installed onto the blotting stack. B. INSTALLATION OF THE BLOTTING STACK 1. Wrap a plastic platform, the size or larger than the gel, with Whatman paper and

2 SOUTHERNS, page 2 place in a plastic container. Fill container 1/3 full with 20x SSC. Allow the Whatman paper to become saturated with the solution. 2. Saturate 3 pieces of Whatman paper a little larger than the size of gel with 20x SSC. Place on top of the platform. Remove air bubbles by rolling a clean test tube over the top. 3. Place gel upside down onto the Whatman paper and remove any trapped air bubbles under the gel by pressing with fingers of your gloved hand. (Gel is placed upside down because the front gel wells are ridged. A flat transfer surface is on the back of the gel.) 4. Cover the gel with a large piece of plastic wrap and with a razor blade cut the plastic wrap around the edge of the gel. Remove the plastic that is on the gel leaving a frame around sides. This ensures that the capillary movement of buffer will only go through the gel. 5. Cut a piece of nylon membrane (Amersham Hybond N) the size of the gel. DO NOT TOUCH THE MEMBRANE WITH YOUR FINGERS! This can cause high background. Use blunt ended forceps and only touch corners!! Wet the nylon membrane with 20x SSC. Carefully lay it on top of the gel, if an air bubble is present lift up on that part of the membrane and lay it down again. Do not touch the membrane or roll a test tube directly on it to remove air bubbles. 6. Saturate 2 pieces of Whatman paper with 20x SSC and place on top of the nylon membrane. Make sure there are no air bubbles by rolling a clean test tube over the top. 7. Cut a stack of paper towels 2 inches thick to fit into the plastic tray and place dry on top of the Whatman paper. 8. Place a flat piece of plastic on top of the blotting stack. Place on top of that a 500g - 1kg weights (e.g. a flask or bottle with water). 9. Allow transfer to proceed overnight. C. TAKING DOWN BLOT 1. With gloved hands remove the layers of the blotting stack down to the nylon membrane. With a pencil, mark the positions of the gel wells on the nylon membrane. 2. Mark the upper left corner of gel by cutting the corner of the nylon membrane off.

3 SOUTHERN,page 3 3 Place the membrane into a tray with 5x SSC for 5 min. Agitate gently. DNA side up. 4. Place the membrane DNA side up onto paper towels. 5. To Bind DNA to the membrane, EITHER place the blot between two pieces of Whatman paper and bake at 80 C in an oven for 2 hours or use the Stratalinker on the Autocrosslink mode on the DNA side and 200 Joules on the other side. 6. Store the dry blot between sheets of filter paper and in a plastic bag at room temp. or for long storage >1 week store at 4 C until ready for the hybridization experiment. Solutions: 1) 0.25N HCl Stock HCl is 12N 20.8ml of HCl 979.2ml of ddh 2 O = 1 Liter 2) Solution D Denature solution 1.5M NaCl 87.75g 0.5M NaOH 20.00g ddh 2 0 to 1 Liter 3) Solution N Neutralizing solution 0.5M Tris 60.50g 1.5M NaCl 87.75g 1mM EDTA 0.372g ddh 2 O to 1 Liter ph to 7.5 4) 20 X SSC 3.0M NaCl g 0.3M NaCitrate 88.2g ddh 2 0 to 1 Liter ph to 7.0

4 SOUTHERN HYBRIDIZATION USING BOEHRINGER MANNHEIM NONRADIOACTIVE NUCLEIC ACID DETECTION SYSTEM A. PREHYBRIDIZATION 1. Place the DNA southern blot into a small hybridization tube by carefully rolling it with the DNA side toward the inside of the tube. Use gloved hands and blunt ended forceps. 2. Warm 20mls of prehybridization solution to 42 0 C. 3. Boil 500ul of Salmon Testes DNA (10mg/ml) for 10min and then place on ice for 2 min. Add the 500ul of DNA to the 20mls of prehybridization solution. The final concentration of DNA is 250ng/ml. This is used to block nonspecific binding sites on the blot. 4. Add the prehybridization solution to the blot in the tube and place tube in hybridization oven that has been warmed to 42 0 C. MAKE SURE YOU BALANCE THE TUBE WITH ANOTHER TUBE OF EQUAL WEIGHT! 5. Prehybridize blot for at least 2hr at 42 o C. Longer times are possible. B. HYBRIDIZATION 1. Prepare hybridization solution for hybridization as follows. Hybridize a 15x15cm blot with at least 6mls of solution and no more than 20mls. The amount varies depending on the amount of stock probe that you have. Warm the Hybridization solution to 42 0 C. 2. Denature Salmon Testes DNA by boiling. (The final concentration of S.T.DNA should be 125 ng/ml in the hybridization solution.) After boiling place on ice 2 min. 3. Denature the DIG-labeled probe DNA by boiling for 10min. (The final concentration of the probe should be 10ng/ml). After boiling, place tube on ice for 2min. 4. Add the probe and Salmon DNA to the warm hybridization solution and mix by shaking well. 5. Remove prehybridization solution from hybridization tube. (The prehybridization

5 Page 2 solution can be reused 2 times. Store at C. To reuse, the Salmon DNA in the solution must be denatured. Place the solution in a 65 0 C water bath for 15min. The flashpoint of pure formamide is 68 0 C therefore do not boil the solution.) 6. Do not let blot dry in tube. Immediately add the hybridization solution and place in hybridization oven. Make sure the tube is balanced! Incubate overnight at 37 o C. C. STRINGENCY WASHES 1. Remove tubes from oven and increase the oven temperature to 65 0 C for future use. Remove hybridization solution. Do not let blot dry in tube. (The hybridization solution can be saved. Store in a 15 or 50ml tube labeled very clearly with the name of probe and # of times used. Reuse 1 or 2 times. To reuse thaw and denature by heating the solution at 65 0 C for 10 min. Cool to hybridizing temperature and add to blot.) 2. Immediately wash membrane twice in the hybridization tube with about 50mls of 2X Wash solution for 15min per wash at room temp. 3. Warm 0.5X Wash solution in 65 0 C water bath and add 50ml to membrane in hybridization tube. Place in hyb. oven which has been warmed to 65 0 C and incubate for 15min. Remove solution and add fresh warmed solution and incubate another 15 min in the oven. 4. Proceed directly to the detection procedure. NOTE: HYBRIDIZATION AND STRINGENCY CONDITIONS: DIG-Labeled probes demonstrate the same hybridization kinetics as radiolabeled probes. Labeled probes can hybridize non-specifically to sequences that bear homology but are not homologous to the probe sequence. These hybrids are less stable than perfectly matched hybrids. They can be dissociated by performing washes of various stringencies. The stringency of washes can be manipulated by varying the salt condition and temperature. It is recommended that one should hybridize stringently, i.e. optimize hybridization conditions (increase temp., but not higher than 55 0 C) rather than washing stringently.

6 Page 3 D. SOLUTIONS 1. Prehybridization and Hybridization solution: 5X SSC 75ml of 20XSSC 2% Block solution 60ml of 10% Block Solution 0.1% N-lauroylsarcosine 3ml of 10% N-lauroylsarcosine 0.2% SDS 6ml of 10% SDS ddh 2 O 6ml of ddh 2 O Formamide (deionized)* 150ml of Pure Formamide 300ml Total Aliquot in 50ml tubes and store at C until use. Salmon Testes DNA and Probes are added just before use. *Deionizing Formamide: Just before preparing the above solution place the desired amount of formamide into a beaker in the fume hood. Add mix resin beads 15g/150ml of formamide to the beaker. Mix with stir bar for 1 hour. Filter the formamide through a piece of Whatman paper. Now it is ready to add to the Pre/Hybridization solution. 2. 2X Wash Solution: 2X SSC 100ml of 20X SSC 0.1% SDS 10ml of 10% SDS ddh ml of ddh Liter 3) 0.5X Wash Solution: 0.5X SSC 25ml of 20X SSC 0.1% SDS 10ml of 10% SDS ddh ml of ddh 2 0

7 Page 4 E. DETECTION 1. After hybridization and stringency washes, rinse membrane briefly with WASHING BUFFER about 2-5 min in a plastic tray with the DNA side up. 2. Incubate membrane in 80mls of BUFFER 2 for 30 min. 3. Dilute anti-dig-ap conjugate 75 mu/ml (1:10,000 dilution) =2ul in 20ml of fresh BUFFER Incubate membrane for 30 min in the antibody solution on shaker either in a plastic tray or in a zip lock bag. Ensure that the solution is covering the entire blot with gentle agitation. 5. In a plastic tray wash the membrane 2X 15 min. with 100ml of WASHING BUFFER. Apply gentle agitation. 6. Equilibrate membrane 2-5 min. in 20 ml BUFFER Dilute CSPD (25mM) a 1:100 dilution in BUFFER 3. A 15X15ml blot will need 2ml of this solution. Add 20ul of CSPD to 1.8ml of Buffer 3. (Note the diluted CSPD can be reused 1X. After use collect the 2mls and place in a 15ml tube, wrap with foil and label. Place at 4 0 C.) 8. Place blot on a piece of Whatman paper to remove excess liquid. DO NOT LET DRY. Using forceps lift and place in a clear plastic sealable folder DNA side up. Pipet the diluted CSPD solution onto the blot. Carefully close the folder. Ensure that the entire blot is covered with the solution. Once closed, immediately seal the folder closed with the sealer. Incubate blot in dark for 5 min. 9. Cut open the folder. With forceps remove blot and place on a piece of Whatman paper to remove excess moisture. DO NOT LET DRY. Keep blot DNA side up. 10. Place the membrane in a new plastic folder and seal closed with the sealer. Incubate for 10 min at 37 0 C. (This enhances the luminescent reaction.) 11. Expose the blot to film. (CSPD is a chemiluminescent substrate for alkaline phosphatase that enables extremely sensitive and fast detection of biomolecules by producing visible light that can be recorded on film. A delay in reaching maximum light emission is observed however the signal persists for several days on the blot.) I usually incubate the blot for 4-6 hours before the initial exposure to film. An exposure of 15min to 1hour should be sufficient however you might have to adjust according to the intensity of signal

8 Page 5 on your blot. CSPD reaches a peak of emission after about 12 hours. Store blot at room temp. in the dark between exposures. Once exposures are completed store blot at room temp. labeled. Blots can be stripped and reprobed. SOLUTIONS: 1) Maleic Acid Buffer 0.1M Maleic acid 11.60g 0.15M NaCl 8.77g ph to 7.5 using NaOH pellets add slowly until ph is reached then add the rest of ddh 2 0 to 1 liter. 2) WASHING BUFFER Maleic Acid Buffer 0.3%Tween 20 Make the same as above 3ml for 1 Liter 3) BLOCKING STOCK SOLUTION 10X conc. Blocking reagent from BMB, 10% (w/v) in maleic acid buffer. Dissolve blocking reagent by constantly stirring on a heating block at (65 0 C). Do not boil the solution. It is difficult to get into solution and may take several hours. Be sure that all of it has dissolved and then autoclave. Store at 4 0 C. The solution is opaque. A much cheaper alternative to BMB blocking reagent is non-fat dry milk. To prepare dissolve 10% (w/v) in maleic acid buffer. DO NOT autoclave this solution. Aliquot and store at C. Short term storage at 4 0 C ( 2-3days). 4) BUFFER 2 (make fresh for each use) 1% Blocking Buffer 10ml Maleic Acid Buffer 90ml Page 6

9 5) BUFFER 3 0.1M Tris-HCl 12.1g 0.1M NaCl 5.84g ddh 2 O 600ml 50mM MgCl 2 25ml of 2M stock_(only needed for color detection not CSPD detection) ph to 9.5 add ddh 2 O to 1 Liter 6) STRIPPING BUFFER 0.1%SDS ddh 2 O 10ml of 10% SDS stock 990ml 1 Liter 7) Anti-digoxigenin (DIG)-AP(alkaline phosphatase conjugate) Fab fragments: Boehringer Mannheim Cat.No , 150 Units 8) CSPD Chemiluninescent Substrate: Boehringer Mannheim Cat.No , 1ml (25mM)

10 NON-RADIOACTIVE DNA PROBE ISOLATION PURIFICATION AND RANDOM PRIMED LABELING USING BOEHRINGER MANNHEIM DIGOXIGENIN SYSTEM A. PROBE SELECTION AND ISOLATION Part 1: Probe Selection and Template Preparation 1. Selection of a probe: The probe should consist of the gene of interest. If possible it should not include any portion of the vector as well as bases outside of the recombination sites. The probe should be greater than 200bp and less than 10kb. 2. Obtaining template DNA: For Random Primed labeling it is most efficient to start with 3ug of template DNA, this should provide enough labeled probe to hybridize > 15 blots or more. The labeling can be scaled up if larger amounts are desired. The following are 2 procedures that can be used to obtain the template DNA. a) If the gene of interest is in a plasmid in which you have PCR primers for and can amplify the gene of interest (e.g. a Bluescript plasmid see below) then amplify 4ug or more (this will give you some to spare) of the gene using the PCR protocols described for that plasmid and primers. After the PCR, test 10ul of reaction on an agarose gel to check for amplification and then quantify your DNA. Proceed to Part 3. Procedure for PCR amplification of genes cloned into Stratagene s Bluescript plasmid called pbs (SK+): 1a. Currently Cry5, PVY and NPTII genes have been cloned into the polylinker site of the pbs (SK+) plasmid vector and the strains are stored in the -80 o freezer. Streak the strain of choice on a LB agar plate containing Ampicillin antibiotic at a concentration of 100ug/ml. Incubate plate at 37 o C O/N. 2a. Inoculate 3mls of LB containing 100ug/ml of Ampicillin with an isolated colony from the O/N plate. Incubate O/N at 37 o C on shaker. 3a. Isolate the plasmid DNA from 1.5ml of the culture using the standard Alkaline Lysis Plasmid Isolation procedure which can be found in Molecular Cloning by Sambrook and Maniatis or by using Promega s Wizard Miniprep Plasmid Isolation System following the instructions given in the kit. When complete quantitate the isolated plasmid.

11 page 2 4a. Setting up the PCR reaction. Two reactions of 100ul volume should produce an amount of DNA in excess of what will be needed for the template in the probe synthesis procedure. In each tube add: Taq Polymerase Buffer 10X dntp mix 10mM MgCl 2 50mM Plasmid DNA T3/T7 primer set from Stratagene Taq polymerase ddh 2 O 100ul 10ul 2ul 3ul 100ng 4ul 0.5ul to 100ul final volume Centrifuge for 5sec to bring solution to bottom of tube. Overlay solution with 30 ul of sterile mineral oil. 5a. Start PCR cycle: 94 o C 5min then 30 cycles of: 94 o C 1min 52 o C 1min 72 o C 2min after cycles: 72 o C 5min 4 o C Holding temp. 6a) Check 10ul of the PCR reaction on an agarose gel to confirm size and quality. If it looks good then determine the concentration. Proceed to Part 3. b) If the gene of interest is not in a plasmid with PCR primers, than you will need to purify at least 100ug of plasmid DNA in order to obtain >3ug of the template DNA. This can be done by using a QIAGEN Midi Plasmid Purification Kit. Qiagen Midi Plasmid Protocol 1b. Growth of bacterial cultures: grow the bacteria in the presence of the selective antibiotic. The quantity of culture will depend on the copy number of the plasmid. Grow enough culture to obtain 100ug of plasmid. Bacterial cultures should always be grown from a single colony grown on a selective plate.

12 page 3 2b. Centrifuge the O/N culture at 5000rpm for 10min. Remove supernatant. 3b. Add RNase A to a concentration of 100ug/ml to the Buffer P1. Add 4ml of this to the bacterial pellet. The bacteria should be resuspended completely, leaving no clumps. 4b. Add 4ml of Buffer P2, mix gently, and incubate at room temperature for 5 min. and not longer. DO NOT vortex, mix by inversion. Vortexing shears the genomic DNA and it will contaminate your sample. Close buffer P2 immediately after use to avoid the reaction of the NaOH with CO 2 in the air. 5b. Prechill Buffer P3 on ice. Add 4ml of P3 and mix gently by inversion immediately. A white precipitate will form. Incubate 15 min on ice. 6b. Centrifuge at > 20,000 x g for 30min at 4 0 C. Remove the supernatant immediately and save. Discard pellet. 7b. Equilibrate a Qiagen-tip 100 by applying 4ml of Buffer QBT and allow the column to empty by gravity flow. The column will not dry out. Do not force out the remaining buffer. 8b. Apply the supernatant to the column and allow it to enter the resin by gravity flow. Once the column is empty, wash the column by applying 2x 10ml of Buffer QC. Allow it to move through the column by gravity flow. Do not force out traces of buffer, it will not effect the elution. 9b. Elute the DNA with 5ml of Buffer QF, allowing it to flow through by gravity. 10b. Precipitate the DNA with 0.7volumes of room temperature isopropanol. Centrifuge immediately at 15,000 x g for 30 min. at 4 0 C. Carefully remove the supernatant. pellets from 2-propanol precipitation have a glassy appearance and may be more difficult to see than the fluffy salt containing pellets with ethanol, therefore mark the outside of the tube before you centrifuge so you can avoid the pellet. 11b. Wash the pellet with 70% ethanol room temperature and redissolve pellet in ddh 2 O. Determine the amount of DNA purified with a spectrophotometer. Proceed to Part 2.

13 Page 4 Solutions For Qiagen: QIAGEN Plasmid Midi Kit (25): includes tip-100 Reagents and Buffers, Qiagen Inc De Soto Ave. Chatsworth, CA Cat. No Provided in Kit: Buffer P1 Buffer QBT Tip-100 column Buffer P2 Buffer QC RNase A Buffer P3 Buffer QF Note: See Qiagen Handbook for recipes for the above solutions if they run out. Part 2: Digestion of Plasmid. 1. Once the plasmid has been purified, proceed with the restriction digestion of the plasmid to produce a fragment of DNA that can be used as a probe. 2. Run a test digest with the enzymes selected on the plasmid and run on an agarose gel to ensure that the correct size fragment appears. 3. Once you have determined which enzymes you will use, digest enough of the plasmid so that you will have at least 4ug of the DNA fragment itself. Part 3: Isolation of Probe DNA Using Qiagen Qiaquick Gel Extraction Kit. Qiagen Qiaquick Gel Extraction Kit: Follow instruction given in kit. B: Random Primed DNA-labeling with Digoxigenin-dUTP. 1. The reagents are available separately or in the GENIUS DIG DNA Labeling Kit from Boeringer Mannheim Cat.No See solutions section for individual Cat.No. 2. The DNA sample should be 500ng-3ug. The 3ug of DNA is most efficient. The sample should be in ddh 2 O at a volume of 15ul as described above in part Heat denature the DNA template in a boiling water bath for 10 minutes, and quickly chill it on ice for 30 seconds before use. Briefly centrifuge tube 10 sec. 4. Add 2ul Hexanucleotide mixture (10X) and 2ul dntp labeling mixture (10X) to the tube on ice.

14 Page 5 5. Add 1ul of Klenow enzyme, labeling grade, to the tube for a final concentration of 100U/ml, and mix using the pipet tip. Incubate the reaction tube at 37 0 C O/N (20Hrs) to obtain 890ng of synthesized DIG labeled DNA. Shorter times will result in less labeled probe. 6. After incubation add 2ul of 0.2M EDTA ph 8.0 to stop the reaction. 7. For all labeling reactions it is extremely important that you verify the labeling efficiency in a direct assay prior to hybridization. Proceed to section C. Solutions for Random Primed DIG-labeling Procedure: DIG DNA Labeling Kit, Random Primed DNA-labeling, from Boeringer Mannheim, Cat. No Hexanucleotide Mix, Cat.No dntp Labeling Mix, Cat.No Klenow Enzyme labeling grade, 100 units, Cat.No Additionally required solutions: 0.2M EDTA ph 8.0

15 Page 6 C: Estimating the Yield of DIG-labeled Probes. 1. Make serial dilutions of the DIG-Labeled control in DNA Dilution buffer according to the following dilution scheme: Labeled Control Stepwise Final Total DNA Dilution Conc.(tube) Dilution 5ng/ul 2ul in 8ul buffer 1ng/ul(A) 1:5 1ng/ul 2ul in 18ul buffer 100pg/ul(B) 1:50 100pg/ul 2ul in 18ul buffer 10pg/ul(C) 1:500 10pg/ul 2ul in 18ul buffer 1pg/ul(D) 1:5000 1pg/ul 2ul in 18ul buffer 0.1pg/ul(E) 1:50000 A->E are the Control dilutions which will be used as standards to quantify your labeling reaction. These samples can be stored at C for continual use. 2. Dilute your probe labeling reaction by making a serial ten-fold dilution. Dilution series for probe labeling reaction: Amount of Dilution Tube Buffer Total Dilution Probe 2ul in 18ul buffer 1:10(A) (A) 2ul in 18ul buffer 1:100(B) (B) 2ul in 18ul buffer 1:1000(C) (C) 2ul in 18ul buffer 1:10000(D) (D) 2ul in 18ul buffer 1:100000(E)

16 Page 7 3. Spot 1ul of each of the dilutions made in step 1 and 2 onto a small piece of nylon membrane, marking the membrane with a pencil to identify each dilution. Mix the dilutions very well just before spotting on the membrane. 4. Fix the nucleic acids to the membrane by crosslinking with the Stratalinker set at autocrosslink. 5. Wet the membrane with 50ml of Washing Buffer. 6. Incubate the membrane in Buffer #2 for 5min at room temperature on shaker with gentle agitation. 7. Dilute anti-dig-alkaline phosphatase antibody 1:5000 in Blocking Solution 10ml. Place membrane in a small zip lock bag with the 10mls of antibody solution. Incubate the membrane for 10 min. on shaker with gentle agitation. The diluted antibody must cover the entire blot. 8. Remove membrane from bag and place in a plastic tray. Wash the membrane 2X 5min in Washing Buffer at room temp. While this is incubating mix 45ul of NBT solution and 35ul X-phosphate solution in 10ml of Detection Buffer. This freshly prepared color substrate solution should be protected from light until use. 9. Incubate the membrane in Detection Buffer #3(with the MgCl 2 for 2 min. Now place the membrane in a zip lock bag and add the diluted color substrate solution to the bag, seal and store in the dark. (Place in a drawer at room temp.) Ensure that the solution is covering the membrane and do not shake it. Let the color development occur in the dark for 30-60min. 10. When the desired spots appear in sufficient intensities, stop the reaction by washing the membrane with Buffer #4 for 5min. Let air dry and store in the dark. The spots will fade in the light. 11. Compare the spot intensities of the control and experimental dilutions to estimate the concentration of the experimental probe. For example, if the spot intensity of the control spot C (10pg labeled control DNA) is equal to the intensity of the D spot of your unknown probe which is diluted 1:1000(=10 3 ) than calculate the amount of DIG-labeled probe DNA to be as follows: 10pg/ul X 10 3 = 10,000pg/ul The total yield of the DNA labeled is the concentration of the DIG labeled probe multiplied by the volume of the probe suspension. If labeling reaction volume was in 20ul than: 10,000 pg/ul x 20ul = 200,000pg or 200ng of labeled DNA Use this as a guide to figure out your concentration.

17 Page Proceed to the Hybridization protocol. Solutions for Estimating Yield of Labeled Probe. DIG Nucleic Acid Detection Kit, Boehringer Mannheim Cat. No includes: Labeled control DNA (Cat. No ) Anti-DIG-AP conjugate (Cat. No ) NBT (Cat. No ) X-Phosphate [BCIP] (Cat. No ) Blocking Reagent (Cat. No ). Use Nonfat Dry instead. It's much less expensive and works the same. These items are available separately see Cat.No. above. Additional solutions: some are the same as the chemiluminescent detection solutions 1) Maleic Acid Buffer 0.1M Maleic acid 11.60g 0.15M NaCl 8.77g ph to 7.5 using NaOH pellets add slowly until ph is reached, then add the rest of ddh 2 0 to 1 liter. 2) WASHING BUFFER Maleic Acid Buffer Tween % Make the same as above 3ml for 1 Liter 3) BLOCKING STOCK SOLUTION 10X conc. Blocking reagent from BMB, 10% (w/v) in maleic acid buffer. Dissolve blocking reagent by constantly stirring on a heating block at (65 0 C). Do not boil the solution. It is difficult to get into solution and may take several hours. Be sure that all of it has dissolved and then autoclave. Store at 4 0 C. The solution is opaque. A much cheaper alternative to BMB blocking reagent is non-fat dry milk. To prepare dissolve 1g/ 100ml of maleic acid buffer. DO NOT autoclave this solution. Short term storage at 4 0 C ( 2-3days).

18 Page 9 4) BUFFER #2 (make fresh for each use) 1% Blocking buffer 10ml Maleic Acid Buffer 90ml 5) DETECTION BUFFER #3 (*MgCl 2 is only needed for color detection not CSPD) 0.1M Tris-HCl 12.1g 0.1M NaCl 5.84g ddh 2 O 600ml *50mM MgCl 2 25ml of 2M stock ph to 9.5 add ddh 2 O to 1 Liter 6) BUFFER #4 10mM Tris-HCl 10ml of 1M ph 8.0 1mM EDTA 2ml of 0.5M ph 8.0 ddh 2 O 988ml of ddh 2 O 1 Liter

19 Page 10 TROUBLESHOOTING Obtain a copy of the Genius System User s Guide for Membrane Hybridization from Boehringer Mannheim. It contains a troubleshooting section that may help you. The biggest problem with the Genius System is background (nonspecific binding to the nylon membrane) which is difficult to strip off. Most often this is due to adding too much probe to the hybridization solution. The procedure involving quantitating the probe can be used as confirmation that your probe has been labeled, however experience has shown that it is sometimes inaccurate for quantitation of the probe. As a guideline the first time I use a newly labeled probe (3ug DNA labeled O/N) I use 1ul of the labeled probe in 20ml of hybridization solution for a 15x15cm blot. This usually works well. If background is present you can: 1) Decrease amount of probe in the hybridization and test a new blot. It will be difficult to strip the blot of the background especially if you are using a DIG labeled standard which limits the stringency of the stripping but you can try it if you want. 2) Increase the amount of DNA in each well to 15->20ug. 3) Do both of the above.