From gilbert@bio.indiana.edu Wed Jan 15 02:45:08 1992 Subject: About Gopher Materials & Methods This is the text content of Jim Brown's nice Hypercard stack of common molecular biology lab protocols. I've indexed it and made it available on the IUBio Gopher server. If you have a useful protocol, please help the community by contributing it to this compendium. Send it via e-mail to jwbrown@crab.bio.indiana.edu For more about this, please read Jim's "GENERAL INFORMATION" >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: GENERAL INFORMATION Materials & Methods (M&M) is a Hypercard stack for the storage & retreival of common laboratory protocols. Each card in the stack contains a single protocol, which can be examined, printed, saved as a text file to disk, or edited. The stack is distributed with a series (lots!) of the protocols which have been used by the author & the Norm Pace lab. New protocols are easily added. SYSTEM REQUIREMENTS This version of M&M (5.1) requires Hypercard version 2.0, or greater. This version of Hypercard requires System 6.0.5 (or greater). Any Macintosh computer capable of running Hypercard 2.0 should run M&M. This means AT LEAST 1Mbyte of RAM, and hopefully a hard disk. M&M cards are the standard 9" size, so that the entire card should be visible on Mac+, SE, or "Classic" screens. You should be familiar with the Macintosh environment in general, and the Hypercard environment in specific, before using M&M. INTRODUCTION M&M stores each protocol on one card, in a scrolling window just like this one. When M&M is started, this "Introduction" card is shown. To browse through the cards, either use the arrow buttons to the right of the card, or click the "Choose card" button. "Choose card" brings up (after an interminable delay) a scrolling list of all of the protocol cards in the stack - just double click on the one you want to go to. To print out a protocol, click on the "Printout" button; to save a protocol as a text (MacWrite) file, usable by any word processing program, just click on the "Save text file" icon. Editing a procedure is no different than in a word processing program - just click at where you want to change something & start typing. Standard cut, copy, paste, delete, etc commands, fonts and font styles, etc, work normally. When you're finished editing, there's NO NEED TO SAVE THE CHANGES TO DISK - Hypercard saves changes automatically & continuously. To create a new protocol card for a new protocol, just select "New card" (command-N) from the Edit menu - you'll then be prompted for a name (keep it short!) for the card, that's used in the "Choose card" list. Shazam! a new card appears with the card name at the top of the text as a reminder. A word of warning - NEVER JUST DELETE ALL OF THE TEXT FROM AN UNWANTED PROTOCOL CARD & RE-USE THE CARD - the name of the card in the "Choose card" list will NOT change, leading to mass confusion. To delete an unwanted protocol card, just select "Delete card" from the Edit menu. After checking to make sure that's really what you want to do, the card is gone. Let me repeat myself here - NEVER RE-USE PROTOCOL CARDS. Importing & exporting text is simple. Click the "Export text" button, & you'll be prompted for a name to save the contents of the current protocol under, as a text (MacWrite) file. To import text, click the "Import text" button & you'll be asked to locate a text (MacWrite or ACSII) file, which will be loaded into a protocol card. If the current protocol card is empty (i.e. just created), the text will go into that card. If the card already contains a protocol, a new card will be generated to contain the contents of the text file (you'll be prompted for a name for the new card, of course). CHANGES IN VERSION 5.1 M&M 5.1 represents a continuation of the trend toward simplification of M&M followed by previous versions. It's been pointed out to me that, like many computer programs, M&M it will probably reach the peak of software perfection at exactly that point in which it ceases entirely to exist. In version 5.1, the "M&M Utilities" stack that came with previous versions is gone - M&M is now (I hope) simple enough not to require it. Also gone are the "M&M" menu, the paging between windows, file transfer, and bugs which prevented correct LaserWriter printing. Version 5.1, unlike previous versions, has "Balloon Help" incorporated into the stack for users of System 7. CONCLUSIONS That's about all there is to M&M. If you have any comments, questions, problems, bug reports, etc, please feel free to contact me at: James W. Brown Department of Biology Indiana university Bloomington, IN 47405 USA or by E-mail at JWBROWN@CRAB.BIO.INDIANA.EDU. I am also happy to give out updates of any of my stacks upon request. This is most easily (for me) done by E-mail, but can also be accomplished by slow-mail. Most of my stuff is also available from the EMBL fileserver. Feel free to copy or give away M&M, on the off chance that anyone should want it, but absolutely positively at NO CHARGE WHATEVER. As you might expect, I cannot guarantee the utility of any protocol that may appear within this stack. Follow any procedure with a critical eye - the person who typed in the protocol was probably in a hurry! If you distribute M&M with protocols you've added, PLEASE be sure to proof-read them! Good luck! Jim Brown >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: EXO III SEQUENCING DELETIONS Reference: Henikoff, S. Unidirectional digestion with exonuclease III creates targeted breakpoints for DNA sequencing. Gene (1984) 351-359. MATERIALS 1. DNA insert cloned into a polylinker containing vector Recommended vector: pBluescript or pTZ. These vectors are specifically designed for Exo III deletions because the restriction sites in the polylinker are arranged so that the 3'-overhang enzyme sites are located on the edges of the polylinker and the 5'-overhang restriction sites are clustered in the interior region. 2. CsCl purified plasmid DNA - miniprep DNA is not pure enough 3. Exonuclease III Recommended enzyme: BRL 8013SA conc. 5000 units in 77 µl 4. Nuclease S1 Recommended enzyme: Boehringer Mannheim 818 330 conc. 400 units/ µl 5. Restriction site selections Do test restriction digests of your plasmid with all restriction endonucleases that are candidates for this Exo III procedure. For deletion in each direction, you need a unique flanking 5'-overhang site and a unique 3'-overhang site which is on the outer edge of the polylinker on the same side of the insert as the 5'-overhanging site. Since exonuclease III requires a 5'-overhang to start deletions from, the sites need to be placed such that the doubly-digested plasmid will be digested into the insert DNA, but the 3'-overhanging cut prevents digestion in the other direction, into the vector DNA. Although the sites must be on the same side of the insert DNA, they should be at least a few bp apart so that both sites will be digested to completion. 6. Exonuclease III Buffer 10 ml from stock solutions 66 mM Tris-Cl pH 8 0.66 ml 1M Tris-Cl pH 8 0.66 mM MgCl2 6.6 ml 1M MgCl2 9.3 ml dH2O 7. S1 Salts Buffer 10 ml from stock solutions 0.25 M NaCl 0.5 ml 5M NaCl 30 mM K Acetate pH 4.6 0.1 ml 3M K acetate pH 4.6 1 mM ZnSO4 .01 ml 1M ZnSO4 5% glycerol 0.5 ml glycerol 8.9 ml dH2O 8. Stop Buffer 10 ml from stock solutions 0.50 M Tris pH 8 5.0 ml 1M Tris pH 8 0.125 M EDTA 2.5 ml 500 mM EDTA 2.5 ml dH2O 9. T4 DNA Ligase New England Biolabs #202 (use at 40 units/µl) 10. Ligase Buffer 10X Buffer: 0.2 M Tris pH 8 0.1 M MgCl2 0.5 M 2-mercaptoethanol 11. Competent cells Recommended strain: DH5a 12. TE : 10 mM Tris pH 8 / 1 mM EDTA EXPERIMENTAL PROCEDURE 1. Doubly restrict 5 µg of plasmid DNA with the chosen restriction endonucleases for one direction. Do a separate restriction digest for the Exo III deletion of your plasmid in the other direction. 2. Phenol extract, add 1/9th volume 3M Na Acetate pH 5.2 and two volumes of 95% EtOH, precipitate on dry-ice EtOH for 10 min. Centrifuge 10 min, wash pellet in 70% EtOH and dry. 3. Resuspend the DNA pellet in 77 µl Exo III Buffer. Mix well and put on ice. 4. Label 35 eppendorf tubes (0.5 ml size tubes) with 30 second interval time points (for each reaction set). Aliquot 25 µl of S1 Salts Buffer into tubes and store on ice. 5. Add 7.6 µl of BRL Exonuclease III (493 units) to the DNA and immediately transfer the tube to a 37¡C waterbath. This is time = 0 min. 6. At intervals of 30 seconds transfer 2.5 µl of the Exo III-plasmid reaction to the S1 Salts Buffer containing tubes, leave on ice. 7. Take time points until all the mixture is gone (about 17 min). A 3 kb insert will be completely digested in approximately 10 min. 8. Prepare an S1 nuclease solution. [1 µl (400 units) in 1103 µl S1 Salts Buffer]. Aliquot 25 µl to each time point, mix and incubate at room temp for 30 min. 9. Add 6 µl Stop Solution to each reaction tube. 10. At this point the time points can be stored at -20¡C (for several weeks, perhaps longer) or prepared for ligation and transformation. Usually it is sufficient to select time points of 1 min intervals (ie. t=1 min through t=10 min) to continue processing. 11. Phenol extract, EtOH ppt, wash and dry selected time points. Resuspend each in 20 µl TE pH 8. 12. Religate the digested plasmids at 14¡C overnight (or 16¡C for 4 hours). Combine: 10 µl Exo III timepoint DNA 2 µl 10X ligase buffer 2 µl 10 mM ATP 1 µl DNA ligase (40 units/ µl) 5 µl dH2O 13. Transformation into competent DH5a cells: Combine: 50 µl competent DH5a cells 10 µl ligation incubate 30 min on ice heat shock for 2 min at 42¡C add 1 ml LB, incubate at 37¡C for 1 hr centrifuge 2 min and resuspend in 100 µl saline or L broth Plate entire transformation on one LB + 100 µg/ml ampicillin plate. Expect 50-300 colonies/plate. 14. Select several colonies from each transformation and start overnight cultures. 15. Prepare minipreps of the cultures and run the samples on a 1% agarose gel. It is important to have two controls on this gel: both your supercoiled original clone and the cloning vector (ideally in the outside wells of both sides of your gel). These controls will serve as markers to size your deletions. Recommended miniprep procedure: Maniatis T. et al (1982) Molecular Cloning: A Laboratory Manual, Alkaline Lysis Method p. 368. 16. Ethidium bromide stain and photograph the gel. Use a ruler to line up your controls and make a list of your deletions in order of increaseing or decreasing sizes. The Exo III deletions may not show perfect progression in the comparison of size to the length of time exposed to the Exo III enzyme. This is not unusual because Exo III is a processive enzyme which will take a single molecule of DNA through a complete digestion before it is released and free to bind another DNA molecule. This should not affect your ability to select clones of all sizes, it may just require screening more colonies to find the appropriate deletion sizes. 17. Sequence the minipreps using the universal and reverse primers and Sequenase 2.0 (see related protocal). Use the primer which is on the same side of the polylinker as your Exo restriction sites. If you use the wrong primer you will sequence the same region of your insert no matter what size deletion you choose (ie. using the vector pBluescript: use the universal primer for Sac I/ Bam HI deletions and the reverse primer for Kpn I/ Sal I deletions). >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: SEQUENCING DOUBLE-STRANDED DNA A variety of sequencing procedures are available - in our hands, the best of these is double-strand sequencing using sequenase and 7-deaza- dGTP. The method is easy and reliable, consistently giving 300 nucleotides of clear, clean readable sequence. Unlike many dsDNA sequencing procedures, supercoiled DNA is used. In fact, miniprep DNA isolated using NaOH-SDS procedures works as well as CsCl-purified DNA. The procedure is, however, quite sensitive to the host-strain of E.coli - DH5alpha works very well, whereas the JM-series work poorly. With CsCl-purified DNA, of course, the host strain doesn't matter. Deaza- dGTP is used to help prevent band compression caused by secondary structure in the DNA product - although not always needed, the sequences are as readable as those using dGTP so we use deaza-dGTP all the time. dITP is not recommended. Sequencing reactions are easiest done 6 at a time (one set of gels worth). SEQUENCING REAGENTS ___________________________________ STOCKS: 10mM dNTPs 10.6 mM 7-deaza-dGTP (USB #70065) 10mM ddNTPs Labeling mix: per ml 10X final conc. _____________________________________________________ 10mM 7-deaza-dGTP 1.5ul 10mM 1.5uM 10mM dCTP 1.5ul 10mM 1.5uM 10mM dTTP 1.5ul 10mM 1.5uM dH2O 995.5ul store 1X & 10X stocks in 100ul aliquots at -20C use 1X stock for sequencing reactions Termination mixes: (final conc: 147uM dATP/dTTP/dCTP/dGTP, 50mM NaCl, 2.7uM ddNTP) solution #1: (standard termination reaction mixes) for 1ml (volume in ul) ddATP ddGTP ddCTP ddTTP ______________________________________________________ 10mM 7-deaza-dGTP 8 8 8 8 10mM dATP 8 8 8 8 10mM dTTP 8 8 8 8 10mM dCTP 8 8 8 8 5M NaCl 10 10 10 10 10mM ddNTP 0.8ddA 0.8ddG 0.8ddC 0.8ddT dH2O 957 957 957 957 solution #2: ("extending mix") for 1ml (volume in ul) _______________________________ 10mM 7-deaza-dGTP 18 10mM dATP 18 10mM dTTP 18 10mM dCTP 18 5M NaCl 10 dH2O 918 combine 2 parts solution #2 to 1 part of each solution #1 store in 100ul aliquots at -20C. Annealing mix (Rx buffer): per ml(1X) final conc _____________________________________________________ 1M Tris-Cl, pH 7.5 200ul 200mM 1M MgCl2 100ul 100mM 5M NaCl 50ul 250mM dH2O 650ul Gel sample buffer per 10ml final conc. ______________________________________________________ formamide 9.5ml 95% 500mM EDTA 0.4ml 20mM bromophenol blue 5mg 0.05% xylene cyanol FF 5mg 0.05% 35S-dATP alpha-35S-thio-dATP at about 1000Ci/mmol, 10uCi/ul Sequencing primers gel or FPLC-purified oligonucleotides at 0.5 pmol/ul (~2.5ng/ul) SEQUENASE SEQUENCING PROCEDURE _____________________ RNase treatment: (skip if samples are already RNased or CsCl) Combine: 2.5ul miniprep DNA (~1ug) 6.5ul ddH2O 1.0ul 100ug/ml RNase A ------------------------------> 10 min. RT (room temperature) Denaturation: Add: 10ul 400mM NaOH ------------------------------> 5 min. RT Add: 3ul 2M NH4OAc pH 4.5 9ul ddH2O 75ul 95% EtOH ------------------------------> 10 min. in dry ice:EtOH bath spin 10 min in microfuge, decant sup fill the tube with -20C 70% EtOH, decant & invert over a paper towel. Tap dry, then dry in speed vac. Sequencing: Add to dried denatured DNA 2ul reaction buffer 8ul 0.5 pmol/ul primer ------------------------------> 10 min. 37C (Use this incubation time to prepare labeling reaction pre-mix and aliquot 3ul termination mixes into each of 4 tubes per reaction.) Prepare labeling reaction pre-mix: 1ul 100mM DTT for 6 tubes: 6.5ul 100mM DTT 2ul 1X labeling mix 13.0ul labeling mix 0.7ul ddH2O 4.6ul ddH2O 1ul 35S-dATP 6.0ul 35S-dATP 0.3ul Sequenase 2.0ul Sequenase Add 5.0ul labeling reaction pre-mix per tube ----------------------------> 2-5 min. RT Termination Reactions: Add 3.5ul of each reaction to 4 tubes containing 3ul of each dideoxy termination mix. ------------------------------> 10 min. 37C Dry in speed-vac Add 10ul gel sample buffer --> vortex (optional: store -20C) SEQUENCING GEL ___________________________________________ Prerun sequencing gels for 30 min at 1200 volts. Heat the samples to 90C for 2 min, and quick-cool on ice before loading. Run 1-2ul aliquots on sequencing gels at 1300-1500 volts (typical gels are 6% polyacrylamide-bis 50% w/v urea) Be careful not to run the gels too hot! For "short" runs, run the xylene cyanol 2/3 to 3/4 of the way down the gel. For long runs; when the xylene cyanol is 3/4 of the way down the gel on the initial loading, load 'blank' sample buffer & run until this second loading of xylene cyanol is 3/4 of the way down the gel. Soak the gel for 15 min. in 10% MeOH, 10% acetic acid, 2% glycerol, lift to 3MM paper & dry in gel drier for 30 min. at 80C. Autoradiograph with the film in DIRECT contact with the dried gel overnight & develop. USB Sequenase protocol USB 7-deaza-dGTP sequencing protocol ESHaas, PaceLabs >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Colony lifts It is very often most convenient to screen clones for the presence of some sequence as colonies, as opposed to isolating plasmid DNA & screening the DNA from Southerns. Although not as "clean" as Southerns, colony lifts are nonetheless a standard lab procedure for screening clones. Although it is possible to lift colonies directly from plated-out transformation reactions, it is best to "grid-out" the colonies first so that extraneous colonies (i.e. blue colonies) are eliminated,allowing more legitimate colonies per plate screened and to make it easier to identify which colony on the plate is giving a hybridization signal. If the gridding is done in duplicate, it also gives you a source of each clone that hasn't been contaminated by the nitrocellulose filter. 1% SDS/5mM EDTA 2X SSC - diluted 1:10 from 20X SSC Lysis solution Neutralization solution 0.5M NaOH 1M Tris, pH8 1.5M NaCl 1.5M NaCl 20X SSC per liter 3M NaCl 175.3 grams 0.3M Na citrate 88.2 grams Adjust to pH 7 with 10N NaOH 1. Spread out some saran-wrap & place 4 3MM squares (large enough for 3 85mm nitrocellulose circles). Pour 1%SDS/5mM EDTA on the first 3MM square, lysis sol'n on the next, neutralization sol'n on the next & 2X SSC on the last. Pour enough solution onto each to saturate the 3MM paper, without leaving pools of liquid on top on the paper. 2. Place a DRY nitrocellulose filter (85mm circle) onto each plate to be lifted & tap down enough so that the filter wets all over. Leave 5 min. 3. Lift the nitrocellulose filters (most of each colony should adhere to the filter) & place colony-side up onto the 1% SDS/5mM EDTA 3MM paper. Incubate 5 min. 4. Transfer the filters to the lysis sol'n 3MM paper & incubate 5 min. 5. Transfer the filters to the neutralization sol'n 3MM paper & incubate 5 min. 6. Transfer the filters to the 2X SSC 3MM paper & incubate 5 min. 7. Place the filters colony-side up onto dry 3MM paper & air-dry 15 min., then bake in a vacuum oven for 2 hr at 80C. The filters are now ready for hybridization as usual. ESHaas >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: CsCl/Et Br plasmid isolation procedure For the purification of supercoiled plasmid DNA which is of maximum purity & is suitable for use with any molecular biological enzymes, the use of CsCl/ethidium bromide is clearly the method of choice. It is, however, relatively expensive in reagent costs (CsCl), materials (disposable centrifuge tubes) and centrifuge time. The time saved working with messy DNA is generally worth it, however. In general, amplification of the plasmid DNA is not required, & is not used in this protocol. STE Lytic mix 100mM NaCl 10% (w/v) sucrose 50mM Tris, pH 8 10mM Tris, pH8 1mM EDTA 5mM EDTA YT Lysozyme sol'n 8g/l tryptone 10mg/ml lysozyme in lytic mix 4g/l yeast extract (prepare fresh) 4g/l NaCl CsCl/EtBr sol'n Triton sol'n 10ml STE 2% triton X-100 10g CsCl - dissolve 50mM Tris, pH 8 for each ml of STE/CsCl, add 80ul 10mg/ml Ethidium Br Saturated 2-PrOH Mix equal volumes of: - isopropyl alcohol - 5M NaCl, 50mM Tris, pH8, 1mM EDTA Shake vigorously, & allow to settle. Use the upper, organic phase. TE 10mM Tris, pH8 1mM EDTA 1. Prepare an overnight culture of the plasmid-bearing strain from which the plasmid will be isolated by inoculating 5-10ml of YT with a drop of a previous culture or a loop-full of cells scrapped from a plate. Incubate overnight at 37C with shaking. 2. Inoculate 1liter YT with 1-2ml of the overnight culture & incubate overnight again at 37C with vigorous shaking. 3. Harvest the cells by low-speed centrifugation (i.e. 5KRPM, 5 min., 4C in an HB4 or GSA rotor) and resuspend the cell pellet in 50ml STE. Repellet 5KRPM, 5 min., 4C in an SS34 rotor and resuspend the cell pellet in 4.5ml ice-cold lytic sol'n. 4. Add 0.9ml lysozyme sol'n, mix in, & incubate on ice for 5 min. 5. Add 1.8ml 250mM EDTA, mix in, & incubate on ice for 5 min. 6. Add 7.2ml Triton sol'n, mix in, & incubate on ice for 20 min. If the suspension does not become viscous, incubate at 37C for 5 min., & recool on ice. 7. Transfer the resulting goo to Ti60 tubes & spin 35KRPM, 1hr, 4C (or use an SS34 at 18KRPM for 1 hr at 4C if you can't get a Ti60). Collect the supernatant (the pellet should resemble an oyster, & have the same rubbery consistency) & measure it's volume. 8. For each ml of cleared lysate, add 1g CsCl. Dissolve gently (don't kick up alot of foam). Remeasure the volume, & for each ml add 80ul 10mg/ml ethidium bromide. 8a. Centrifuge 10KRPM (SS34 or HB4 rotor in Sorval) in Corex tubes. Recover the CsCl/EtBr/extract solution from underneath the EtBr/protein/junk pellicle. 9. Divide into tubes for the Ti75, Ti60, VTi60 or VTi65, top off with CsCl/EtBr sol'n (or add 3.5ml @ to SW50 rotor tubes, fill & balance with mineral oil) & close the tubes (except SW rotors) with caps or heat-sealer. Load the tubes into the rotor & centrifuge as specified: VTi60, VTi65 ON 20C 45KRPM Ti75, Ti60 2days 20C 40KRPM SW50, SW50.1 ON 20C 35KRPM 10. Using a short-wave UV light to visualize the DNA bands, collect the lower, plasmid DNA bands by syringe through the sides of the tubes (don't forget to puncture the top of the tube if its heat-sealed, or open if it's capped, before inserting the syringe). If you have lots of DNA, the band should be easily visible without need for UV illumination. 11. If using a vertical rotor, pool the samples & transfer to a fresh centrifuge tube. Top off with CsCl/EtBr sol'n & centrifuge as before. Once again, collect the plasmid DNA band via syringe through the side of the tube. 12. Mix with an approximately equal volume of Sat'd 2-PrOH, & allow to settle for a few minutes. Collect & discard the upper, 2-PrOH phase. Repeat this until both phases are colorless, then repeat twice more for good measure. 13. Dialyse overnight with 2 changes of TE at 4¡C. 14. Store at 4C. Examine the DNA on a minigel & extimate the concentration either by UV spectrophotometry (A260) or using an EtBr spot test. Maniatis, Mol. Cloning Manual Davis, Mol Cloning Manual Davis Cram, personal communication Greg Beckler, personal communication >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Isolation of total RNA using CsCl/EtBr gradients RNA for S1 or PE analysis must be free of chromosomal DNA. This can be accomplished by extraction of the RNA with hot phenol, but phenol can, especially if its pH is acidic, cause the specific loss of certain RNAS, i.e.polyadenylated RNAs. This method, which is simply lysis by Frech pressing in RNase inhibitor-containing buffer followed by phenol:chloroform extraction, EtOH precipitation & CsCl/EtBr isopycnic centrifugation, is relatively rapid, easy, & results in an undegraded RNA preparation essentially free of DNA (without the use of DNase). Because the procedure does not rely on cell wall hydrolytic enzymes (lysozyme) or any other specific properties cell envelop, RNA from almost any type of cell can be isolated. Remember to use RNase-free technique throughout this procedure. Use DEPC-treated ddH2O, & absolute EtOH. And remeber - Ethidium bromide is MUTAGENIC! Wear gloves & clean up after yourself. A Ti50 or Ti75 rotor can also be used - more tubes will, of course, be required. STE-SDS 100mM NaCl 50mM Tris, pH 8 1mM EDTA 0.1% SDS CsCl/EtBr sol'n 10ml STE-SDS 10g CsCl - dissolve for each ml of STE/CsCl, add 80ul 10mg/ml Ethidium Br Saturated 2-PrOH Mix equal volumes of: - isopropyl alcohol - 5M NaCl, 50mM Tris, pH8, 1mM EDTA Shake vigorously, & allow to settle. Use the upper, organic phase. SSI Lysis buffer (per 200ml) final conc. working conc. 1.9380 grams Tris,pH 7.5 80 mM 35.6 mM 0.4067 grams MgCl2-7H2O 10 mM 4.4 mM 140 ul 2-mercaptoethanol 10 mM 4.4 mM SSI lytic mix (per 200ml) 2g SDS 1% 0.5% 40ml 0.1M EDTA 20mM 10mM 158mg 1,10-phenanthroline 4mM 2mM 160mg heparin 400ug/ml 200ug/ml SSI mix is prepared by mixing equal volumes of SSI lysis buffer & SSI lytic mix. 1. Grow 2liters, to mid-log phase, of strain from which RNA will be isolated. Harvest by centrifugation at 5KRPM, 4C 10 min. (GSA or GS3 rotor). 2. Resuspend the cells in 20ml SSI mix & French press at 20000 PSI. 3. Mix the lysate with 20ml Phenol:chloroform. Vortex & centrifuge 10KRPM, 4C 10 min. (HB4 or SS34 rotor), then collet the upper, aqueous, phase by pipette. 4. Repeat step 3 twice more. 5. Add 1/10th volume of 3M Na acetate and 2 volumes of EtOH. Incubate for 1hr at -70, then centrifuge at 10KRPM at 4C for 20 min. (SS34 or HB4 rotor). Discard the liquid & drain the pellets dry of EtOH. 6. Dissolve the RNA in 20ml STE-SDS, then carefully remeasure the volume of the mixture. 0.11 volumes of 2mg/ml ethidium bromide, then add 1g of CsCl for each ml of the RNA/EtBr solution, and thoroughly dissolve. 7. Load the solution into a sealable Ti60 tube. Top off the tube with blank CsCl/EtBr sol'n, & seal. Load the tube (and a balance) into the Ti60 rotor and centrifuge for 2 days at 38K, 20C. 8. Collect the DNA band via syringe through the tube (don't forget to pucture the top of the tube first) if desired. Cut the tube in half, and drain the CsCl/EtBr sol'n off of the RNA pellet. 9. Dissolve the RNA (it may take some work) in ddH2O, then add 1g of CsCl for each ml of RNA sol'n. 10. Add an approximately equal volume of Sat'd 2-PrOH, mix, & allow to resettle. Collect the upper phase & discard. Repeat until no pink color remains in either phase, then repeat twice more to be sure all of the ethidium bromide is gone. 11. For each ml of CsCl/RNA sol'n, add 2ml of H2O and 9ml of EtOH. Incubate overnight at -20C. 12. Centrifuge 20min, 10KRPM 4C (HB4 or SS34 rotor), & discard the supernatant. Drain the RNA pellet dry & dissolve in 1ml ddH2O. 13. Remove a small aliquot (20ul) for analysis in a minigel (2% agarose). Store the RNA at -20C. JWBrown, unpublished data Maniatis "Molecular Cloning" manual >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: DMS-treating DNA DMS treatment under controlled conditions, followed by piperidine cleavage, yeilds a ladder of fragments indicating the positions of all G residues in the DNA sequence. These ladders are useful MW standards for S1 experiments, footprinting, etc. DMS buffer Sequencing dye --------------- ---------------- 50mM Na Cacodylate,pH8 80% formamide 1mM EDTA 10mM NaOH 1mM EDTA DMS STOP 0.1% xylene cyanol FF -------------- 0.1% bromophenol blue 1.5M NaOAc 1M 2-ME Carrier DNA ---------------- 1mg/ml sonicated or sheared salmon sperm DNA 1. Dry the desired amount of singley end-labeled DNA in an eppendorf tube (you need 5000-20,000 CPM per lane on a gel). 2. Resuspend the DNA in 200ul ice-cold DMS buffer. Working on ice, add 3ul carrier DNA, vortex, and add 1ul DMS. Vortex again, and incubate 5 min on ice. 3. Add 50ul DMS STOP and 750ul EtOH, mix, and freeze for 5 min in a dry- ice:EtOH bath. Spin for 5 min and discard the supernatant. 4. Dissolve the pellet in 250ul 0.3M NaOAc, then add 750 EtOH and freeze & spin as before. Add 1ml ice-cold 70% EtOH to the pellet, and freeze & spin again. Discard the supernarant and dry the pellet in a roto-vac. Add 20ul ddH2O to the dry pellet and dry again. 5. Add 100ul 1:10 piperidine (freshly diluted in ddH2O) and incubate for 30min at 90C. Quick-freeze the solution in a dry-ice:EtOH bath, then dry the sample in a roto-vac (with the heat ON to speed the drying). 6. Add 10ul ddH2O to the dry pellet, and redry. Repeat this step. 7. Count the pellet and resuspend the DNA in enough sequencing dye for 3-5ul per lane. Remember to heat the sample for 5min at 90C, then cool on ice before loading the gel. Meth Enzymology 65, Maxam & Gilbert >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: DNase I footprinting DNase I footprinting is a method for determining the site of binding for a protein on a DNA sequence. In order to use this procedure, you must have: purified protein, the DNA sequence of the cloned DNA in question, and filter-binding data indicating the general region of binding (or at least good reason to believe the binding site is within a relatively small region). Before starting the actual DNase I footprinting, you need to isolate singley end-labeled probes of both strands containing the binding region, dissolved in ddH2O to 10,000 - 20,000 CPM/ul. These probes should be tested by DMS reactions (use 1/10 of the total amount of each probe - the rest of the DMS-treated probe is used for MW markers during the footprinting) for purity and to be sure the probes are the correct fragments. Once you have these probes, you must determine the correct amount of DNase I required for apprpriate partial digestion of EACH PROBE empirically by titration. With this information you are ready to begin DNase I footprint assays. Both the titration & footprinting are explained here. DNase I stock - 1mg/ml RNase-free DNase I in TMK buffer Store at -20C. TMK buffer 2X footprint buffer ------------- --------------------- 50% glycerol 100mM KCl (adjust to optimize binding) 50mM Tris pH8 20mM MgCl2 50mM KCl 40mM Tris pH8 10mM MgCl2 0.2mM EDTA NTP mix STOP solution ----------- --------------- 3.3mM ATP 0.1M EDTA 3.3mM GTP 0.6M NH4OAc 3.3mM CTP 20ug/ml sssDNA DNase I titration 1 - For each probe, mix (working on ice): 440ul ddH2O 50ul 5mg/ml BSA (nuclease-free) 500ul 2X footprinting buffer 50ul NTP mix 10ul probe DNA Divide the mix into 10 reactions of 100ul each, in eppendorf tubes. 3 - Dilute 40ul stock DNase I in 116ul TMK (=256ug/ml DNase I). Do 8 1:2 serial dilutions of this 256ug/ml solution (75ul TMK + 75ul DNase I, mix, go to the next dilution). 4 - With all samples starting on ice, the experimental time course is : tube # 1 2 3 4 5 6 7 8 9 10 -------------------------------------------------------- --> 30C 0min 1 2 3 4 5 6 7 8 9 DNase(5ul) 10 11 12 13 14 15 16 17 18 19 100uLSTOP->ice 12 13 14 15 16 17 18 19 20 21 -------------------------------------------------------- [DNase] used 0 1 2 4 8 16 32 64 128 256 5 - When all the samples are done, add 100ul phenol:chloroform to each & vortex. Spin 1-2 min and transfer the upper phases to tubes containing 600ul EtOH. Freeze the samples in a dry-ice:EtOH bath for 5 min, then spin for 5 min. Discard the supernatants and add 750ul ice-cold 70% EtOH to each tube, refreeze and spin as before. 6 - Resuspend each sample in 5ul sequencing dye and run a 6% sequencing gel. Run an appropriate distance for the size of the DNA used (bromophenol blue to bottom for 150bp, XC to bottom for 400bp). Dry & autoradiograph the gel (overnight exposure with screens at -70C). Choose the best partial digest for future use - you want even cleavage at all sizes (not faded at the top) with uncut DNA left over and the darkest partial bands. Footprinting 1 - Working on ice, mix the same reaction mix as before for each probe. Divide the mix into 10 tubes each of 100ul. Prepare dilutions of DNase I from the stock as before. You will use 5 different dilutions - the optimum and 2-fold dilutions up and down from this optimum (i.e. if 1:32 is best, use 1:128, 1:64, 1:32, 1:16, & 1:8) to make sure you get the best results. 2 - With all samples starting on ice, the time schedule is : Tube # 1 2 3 4 5 6 7 8 9 10 --------------------------------------------------------- 1ulprotein->30C 0 1 2 3 4 5 6 7 8 9 +5ul DNase I 10 11 12 13 14 15 16 17 18 19 100ulSTOP->ice 12 13 14 15 16 17 18 19 20 21 --------------------------------------------------------- [DNase] used .25 .5 1X 2X 4X .25 .5 1X 2X 4X relative to opt. protein added? - - - - - + + + + + --------------------------------------------------------- If required by protein concentration, more (up to about 10ul depending on what it's dissolved in) binding protein can be used. 3 - Phenol:chloroform extract, EtOH precipitate, 70% wash, dry, resuspend & run the 6% sequencing gel as before. Dry the gel and expose as indicated by experience from the DNase I titration. Repeat the footprinting using ddH2O in place of NTP mix. Examine the autoradiographs for gaps in the DNase I ladder found in the samples with added binding protein but not in the lanes without added RNAP. JWB, unpublished Craig & Nash 1984 Cell 39:707-719 Galas & Schmitz 1978 Nucl. Acids Res. 5(9):3157 >>> From jwbrown@crab Wed Jan 15 02:45:08 1992/ Subject: dscDNA synthesis The method described here is for the synthesis of blunt-ended double-stranded cDNA, suitable for cloning either by blunt-end ligation or linker-ligation into a plasmid or M13 vector. The most important factor in synthesizing cDNA is that the mRNA template be as pure as possible - free of RNase, rRNA, inhibitory compounds (especially polysaccharides), and, if a specific cDNA is desired, extraineous poly(A)-containing RNAs. This is a technically difficult technique, so don't be discouraged if it doesn't work the first couple of times. If it doesn't work, retest the poly(A)-containing RNA by agarose-urea electrophoresis to see if it is still intact. Strict RNase-free technique should be used throughout this procedure. All glassware, plasticware, and buffers should be autoclaved, except for the dNTPs, 2-ME, and oligo(dT)12-18. NOTES * If 32P and 3H is needed only to trace the reaction, the isotopes can be decreased to 5uCi each. * If you desire or need to use an RNase inhibitor, then replace the 2- ME with 2.5ul 0.1M DTT and add 5 units RNasin and 100ug/ml BSA during the first strand synthesis. 10X cDNA buffer (per 100ml) final conc. --------------- -------- 6.06 grams Tris-OH 0.5M Adjust to pH8.3 2.03 grams MgCl2-7H2O 0.1M at 42C with 10.44 grams KCl 1.4M conc. HCl 0.3M 2-mercaptoethanol - 131ul 2-ME plus 4.87ml ddH2O 20mM dNTPs dATP - 25mg plus 2.15ml ddH2O dCTP - 25mg plus 2.35ml ddH2O dTTP - 25mg plus 2.23ml ddH2O dGTP - 25mg plus 2.12ml ddH2O Add ddH2O directly to the bottle, dissolve, and adjust to neutrality with 50mM Tris, pH7 dNTP mix - mix 5ul of each 20mM dNTP just before starting. TEN (per liter) 1.2114 grams Tris-OH, pH 7.8 10mM 0.5844 grams NaCl 10mM 0.3802 grams Na4EDTA 1mM 8u/ml oligo(dT)12-18 - Add 125ul ddH2O/A260 unit directly to the vial 1M HEPES, pH 6.9 - 5.206 grams per 20ml S1 nuclease buffer (per 100ml) 0.4082 grams Na acetate 30mM 1.7532 grams NaCl 0.3M 0.0409 grams ZnCl2 3mM 2.5M Na acetate - 34.02 grams per 100ml 25:1 chloroform/1-octanol - 10ml 1-octanol + 240ml chloroform First strand final reaction mixture: 50mM Tris, pH 8.3 (at 42C) 10mM MgCl2 140mM KCl 30mM 2-ME 0.04 A260 units oligo(dT)12-18 1mM @ dATP, dCTP, dGTP, and dTTP 50uCi 32P-dCTP 5ug poly(A)-containing RNA 50 units reverse transcriptase ------------------------------ 50ul total volume Second strand final reaction mixture: 0.1M HEPES, pH 6.9 0.75mM @ dATP,dCTP, dGTP, and dTTP 50uCi @ 3H-dCTP and 32P-dCTP 70mMKCl 100 units DNA polymerase I 5mM MgCl2 15mM 2-ME other sscDNA syn. leftover --------------------------- 100ul total volume First strand synthesis: 1. Transfer 50uCi a32P-dCTP to a 500ul microfuge tube, plug it with cotton, and cover it with a double layer of parafilm. Puncture the parafilm several times with a needle. 2. Dry in a vacuum dessicator under vacuum to complete dryness (usually 15-30 min.). Remove and discard the parafilm and cotton. 3. Working on ice, add the following reagents to the vial: 5ul 10X cDNA buffer 5ul 0.3M 2-ME 5ul 8u/ml oligo(dT)12-18 10ul dNTP mix 5ul 1ug/ul poly(A)-containing RNA 12.75ul ddH2O 4. Vortex, and add 7.25ul reverse transcriptase (50 units). GENTLY MIX, spin 5 sec., and incubate for 1 hr at 42C. 5. Drop in a boiling water bath for 2 min., and rapidly cool on ice. Remove a 2.5ul aliquot for later assaying. Second strand synthesis: 6. Centrifuge the remainder for 1 min., and transfer the liquid (47ul) to a 500ul microfuge tube containing 5uCi 3H-dCTP dried down as described in steps 1 & 2. 7. Add 10ul 1M HEPES, pH 6.9, 5ul dNTP mix, and 34ul ddH2O to the vial, and vortex. Then add 13ul DNA polymerase I (100 units), gently mix again, and incubate at 15C for 3 hr. 8. While the reaction mix is incubating, assay the sscDNA synthesis as follows: 1. Dilute a 2.5ul aliquot of sscDNA with 40ul TEN. 2. Add 5ul glycerol, and vortex. 3. Apply this sample to a 0.5 x 30cm sephadex G50(50- 150) column, equilibrated with TEN. 4. Collect 32 @ 25 drop (0.5ml) fractions into 6ml scintillation vials containing 5ml scintillation fluid each. 5. Cap and mix the vials, and count them by scintillation spectrophotometry. 9. After incubating, add 10ul glycerol and 110ul 25:1 chloroform/1- octanol, and vortex. Centrifuge 1 min. to separate the phases. 10. Using a pipette gun, collect the upper (aqueous) phase, being very careful not to get any of the lower (organic) phase. Apply the entire aqueous phase to a 0.5 x 30 cm Sephadex G50(50-150) column (equilibrated and run in TEN.). 11. Run the column, and collect 36 @ 25 drop (0.5ml) fractions. Transfer a 25ul aliquot from each fraction into separate 6ml scintillation vials containing 5ml of scintillation fluid. Count each fraction using separate 32P and 3H windows. 12. Pool the fractions containing the cDNA peak, and add 1/10 volume of 2.5M Na acetate and 2 volumes of 2-PrOH. Store overnight at -70C. S1 nuclease treatment: 13. Pellet the dscDNA by centrifuging for 1 hr at 35KRPM (SW55Ti rotor) at 4C. Discard the supernatant, allow the excess fluid to drain off, and dry the pellet under vacuum. 14. Dissolve the invisible pellet by adding 520ul S1 nuclease buffer and a baby stir bar (autoclaved). Stir very slowly overnight at 4C. 15. Transfer the solution to a 1.5ml microfuge tube, and spin 1 min. to remove junk. 16. Transfer to a fresh 1.5ml microfuge tube, then remove a 25ul 'Pre-S1 nuclease' aliquot for later analysis (store at -20C). Add 6ul S1 nuclease (60 units) and incubate at 37C for 1 hr. 17. Remove a 25ul 'Post-S1 nuclease' aliquot for later analysis (store at -20C). Add 500ul 25:1 chloroform/1-octanol and vortex. 18. Centrifuge for 1 min. to separate the phases. Carefully collect the upper (aqueous) phase, and transfer it to a fresh 1.5ml microfuge tube. Add 1ml 2-PrOH, mix, and store overnight at -70C. 19. Pellet the dscDNA for 1 hr, and decant the supernatant. Allow the pellet to drain dry, the dry under vacuum. Store at 4C until use. 20. Treat the pre- and post- S1 nuclease samples exactly as described in step 2, but count the 3H and 32P channels separately. Rosemarie Spencer (personal communication) Buell, et al. 1978 J. Biol. Chem. 253:2471-2482 Wickens, et al. 1978 J. Biol. Chem. 253:2483-2495 Land, et al. 1981 Nucl. Acids Res. 9(10):2251 J. Vaughn (personal communication) BRL Product Profile: AMV Reverse transcriptase Monahan, et al. 1976 Biochemistry 15:223-233 Baulcombe and Verma 1978 Nucl. Acids Res. 5(11):4141 >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: End-labeling DNA fragments End-labeled DNA fragments are used in a wide variety of molecular biology experiments. Doubley end-labeled fragments are useful as MW standards on Southern blots, as probes for filter-binding or gel retardation experiments, or for visualizing small amounts of restricted DNA. Singley end-labeled DNA is used for Maxam & Gilbert sequencing, for S1 analysis, footprinting, primer extension, etc. This method can also be used for 5' end-labeling RNA molecules. 10X Kinase buffer Prep Gel ------------------ ---------------- 0.5M Tris,pH9.5 48.3ml ddH2O 0.1M MgCl2 25ml glycerol 50mM DTT 60mg APS 50% glycerol 16.6ml 29%:1% acrylamide:bis 10ml 10X TBE Denaturation buffer -------------------- 1M Tris, pH9.5 Elution buffer 10mM Spermidine ---------------- 1mM EDTA 0.5M NH4OAc 10mM Mg(OAc)2 Phenol:chloroform 1mM EDTA ------------------- 0.1% SDS 48% chloroform 50% TE sat'd phenol 2% isoamyl alcohol 1 - Digest 10ug DNA with the appropriate restriction enzme (the one you want to label at) overnight in a 30ul reaction. If you desire or need to, a larger reaction can be used, then the DNA should be EtOH precipitated and resuspended in 30ul 1X restriction buffer. 2 - Add 3.5ul 1M Tris, pH8, and 2ul 4u/ul CIAP (calf intestine alkaline phosphatase). Incubate 30 min at 56C. 3 - Add 56ul 2M NH4OAc and 156ul ddH2O. Add 200ul phenol:chloroform, vortex VIGOROUSLY, then spin 1 min. Collect the upper (aqueous) phase and again add 200ul phenol:chloroform, vortex, and spin as before. Collect the upper phase, and add 600ul EtOH and freeze for 5 min in a dry-ice:EtOH bath. Spin for 5 min, discard the supernatant, and add 750ul ice-cold 70% EtOH. Again freeze for 5 min in a dry-ice:EtOH bath, spin for 5 min, and discard the supernatant. Dry the pellet in a rotovac. 4 - Dissolve the pellet (likely invisible) in 20ul ddH2O, then add 20ul 2X denaturation buffer. Incubate for 15 min at 65C (95C for 3' overhangs), and quick-chill on ice. 5 - Add : 6ul 10X kinase buffer, 100uCi gamma-32P-ATP, ddH2O to 59ul, and 20u polynucleotide kinase. Incubate for 1hr at 37C. 6 - Add 56ul 2M NH4OAc and 134ul ddH2O. Phenol:chloroform extract, EtOH precipitate, wash, and dry as in step 3. --- for singly end-labeled fragments --- 7 - Dissolve the dry DNA in 40ul ddH2O, ad 5ul 10X restriction enzyme buffer,and 5ul of the appropriate restriction enzyme. Incubate at the appropriate temperature for 3-5hr. 8 - Add 10ul 10X tracking dye and load onto a 2mm thick 5% prep gel. Electrophorese overnight at 160-180V (for our standard 23cm length gels) in TBE. 9 - Remove one glass plate and cover the gel with saran-wrap. In the dark room, cover the gel with a small piece of X-ray film for 2 min. Develop the film. When the film is dry, cut out the exposed area for the bands you want to elute, then place the film over the gel PRECISELY as it was exposed (use the tracking dye & wells as guides) and mark the saran-wrap with a sharpy where you've cut out the image of the bands. Remove the film, then cut the DNA containing regions from the gel with a razor blade. Dice the gel slice with the razor blade. 10 - Flame-seal a blue 1ml pipette tip & plug the tip with a small amount of glass wool. Put the diced acrylamide in the tip, & wash it to the bottom with 500ul elution buffer. Cover the top with parafilm, vortex, and incubate overnight at 37-42C. 11 - Carefully puncture the sealed-up tip with a red-hot needle, then put the tip point-down into a 4ml snap-cap tube with the top half of the lid in place (cut off the bottom of the lid with a razor blade, leaving a ring that snaps onto the top of the tube). Puncture the parafilm with a red-hot needle, and spin for 1 min in a clinical centrifuge. Add 100ul elution buffer to the tip and spin again. Collectthe eluted DNA from the snap-cap tube and transfer to and eppendorf tube. 12 - Add 1ml EtOH and freeze for 5 min in a dry-ice:EtOH bath, spin for 5 min, and discard the supernatant. Wash 3 times by adding 750ul ice- cold 70% EtOH, freezing, and spinning as before. Dry the final pellet in a roto-vac, and count in a scintillation counter. Elio Vanin, OSU Dept. Biochemistry Maxam and Gilbert, Meth. Enzymol. >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: End-labeling RNA RNA samples can be 5' or 3' end-labeled using commercially available materials. 3' end-labeling is carried out by RNA ligase, which 'ligates' a single labeled nucleotide to the 3'-OH end of the RNA. The labeled substrate in a 3',5'-bis-phosphate (usually cytidine 5',3'- bis-phosphate, 32P-pCp), so that the resulting 3' end is a phosphate, and no longer a substrate for the enzyme. 5' end-labeling is carried out exactly as for DNA (see that procedure in M&M as well), unless the 5' ends are capped; in that case, the RNA must first be de-capped with tobacco acid pyrophosphatase (TAP). Naturally, RNase-free technique MUST be used at all times, including wearing gloves at all times, using sterile autoclaved or DEPC treated materials and high-purity chemicals. MATERIALS FOR 3' END LABELING 10mM Tris-HCl, pH7.4 5X RNA ligase buffer 250mM HEPES, pH8.3 50mM MgCl2 16.5mM DTT 0.5mM ATP 10X GE tracking dye (as for glyoxal gels) 10mM Tris-HCl, pH7.6 Materials of 5' end-labeling 10X TAP buffer 0.5M Na acetate, pH6 0.1M 2-mercaptoethanol 0.5M Tris-HCl, pH8.3 at 37C 250mM KH2PO4, pH9.5 (with KOH) 250mM MgCl2 50mM dithiothreitol 10mM Tris-HCl, pH7.6 PROCEDURE FOR 3' END-LABELING 1. Working on ice, mix the following: 6ul 5X RNA ligase buffer 3ul DMSO 3.5ul glycerol 5ul (50uCi)32P-pCp 1ul (0.5-1.0ug) RNA 9ul ddH2O 2.5ul (10 units) RNA ligase --------------------------------- 30ul 2. Incubate for 24hr at 4C, then stop the reaction at 65C for 5 min. 3. Add 60ul 10mM Tris and 10ul 10X GE tracking dye. Apply the sample to a 6x150mm G-50 sephadex column, in 10mM Tris. Collect 35 12drop fractions. 4. Spot 5ul from each fraction to a filter paper square, and count in 5ml Aquasol. 5. Pool the void volume peak CPM fractions, and store frozen. PROCEDURE FOR 5' END-LABELING CAPPED RNAs 1. Mix the following ingredients, on ice: 1ul 10X TAP buffer Xul RNA (up to 1ug) Yul ddH2O 1ul TAP (2-5 units) ------------------------ 10ul final volume 2. Incubate 37C for 30 min, then add: 2ul 0.5M Tris-HCl, pH8.3 7ul ddH2O 1ul CIAP (or BAP)(4 units) 3. Incubate at 56C (or 37C for BAP) for 30 min, then add 1ul K-PO4, pH9.5 & mix. 4. Transfer the mixture to a fresh tube containing 20-50uCi gamma32P- ATP dried down in the tube. Add 1ul 250mM MgCl2, 2ul 50mM DTT and 1ul (4-10 units) polynucleotide kinase. 5. Incubate 37C for 30 min. Add 5ul 10X GE tracking dye and 20ul 10mM Tris-HCl, pH7.6. Apply the sample to a 6x150mm G-50 column (in 10mM Tris, pH7.6), and collect 35 12drop fractions. Spot 2ul samples of each fraction onto filter paper squares and count them in 5ml Aquasol. 6. Pool the void volume peak CPM fractions and store frozen. NOTE: For use with non-capped RNAs, mix the RNA with 2ul 0.5M Tris-HCl, pH7.6 + 0.1M 2-ME, 1ul CIAP and ddH2O to 20ul, then start at step 3. England 1978 Nature 275:560 Brown 1985 J.Bacteriol. in press Efstratiadis 1977 NAR 4:4165 >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Ethidium bromide spot test for DNA or RNA concentration estimation The concentration of DNA or RNA in a sample is most accuratly determined by absorbance at 260nm. More often, the sample is too small or too dilute for this method, which generally requires ug amounts of the sample for accurate measurement. A fast and relatively accurate method for RNA/DNA concentration estimation is the EtBr spot test. This method will usually allow the estimation of concentration using only ng amounts of sample. 0.1ug/ml standards - Dissolve lambda DNA or E.coli rRNA in TE to ~1ug/ml, then accurately determine the concentration by A260 (an A260 of 1 = 50ug DNA/ml or 40ug RNA/ml). Dilute to 100ng/ml in TE, & preparae dilutions of 90, 80, 70, 60, 50, 40, 30, 20, and 10ng/ml from this stock. 1. On a petri dish or glass slide, spot 10ul aliquots of 10, 20, 30, 40, 60, 80 and 100ng/ml ribosomal RNA or lambda DNA (depending on whether your sample is RNA or DNA). 2. Prepare serial 1:2 dilutions of your sample (usually 5-8 are enough) in TE, and spot 10ul of each of hese dilutions onto the petri plate. 3. To each spot, add 10ul 0.2ug/ml ethidium bromide, & mix by pipetting up & down. 4. Using a transilluminator or hand-held UV source, photograph the plate. 5. Estimate the concetration of the sample by conparison of the dilutions of the samples with each of the standard DNA or RNA samples. >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Eubacterial RNA isolation A variety of RNA isolation procedures are available for different species of eubacteria - it is useful, however, to have a "general" method for isolating RNA from any specie you start to work with, such as the procedure described here. This procedure generally yeilds 20-50mg of RNA at good purity. No effort is made in this procedure to eliminate DNA, which is a minor contaminant. This procedure is primarily useful to those working with stable RNAs, or others needing large quautities of RNA. Rapidly turned-over RNAs should be isolated from RNA isolated by a more rapid procedure, such as the single-step RNA isolation procedure of Gopalakrishna, et al. STE-SDS (for STE, omit SDS) 100mM NaCl 10mM Tris, pH8 1mM EDTA 0.1% SDS Lytic solution Triton solution 10% sucrose 2% Triton X-100 10mM Tris, pH8 50mM Tris, pH8 5mM EDTA Lysozyme solution - 10mg/ml lysozyme in Lytic solution 250mM EDTA 3M NaOAc (sodium acetate) Sat'd phenol - add an equal volume of 1M Tris (pH8) to some phenol, shake well, then allow to settle. Discard the upper phase (aqueous) & add an equal volume of TE. Mix well, & store at 4C. 1. Grow up enough of the organism to late log to get about 20g wet weight of cell paste (try 4 liters). (The procedure can be used with less paste by scaling down.) Cool the cells on ice, then pellet by centrifugation at 5KRPM , 10min., 4C (GS3 or GSA rotor), wash in 100ml STE or saline and recentrifuge. Discard the supernatants. 2. Resuspend the cells in 20ml lytic solution, add 4ml lysozyme sol'n, mix, & incubate 5 min. on ice. 3. Add 8ml 250mM EDTA, mix, & incubate on ice 5 min. 4. Add 30ml triton solution, mix, & incubate 20 min. on ice. 5. French press the suspension at 20KPSI, and remove cell debris by centrifuging at 10KRPM, 4C,10min HB4 rotor. Collect the supernatant. 6. Add an equal volume of sat'd phenol, mix vigorously, and centrifuge 10KRPM, 4C, 10min (HB4). Collect the upper (aqueous) phase. Repeat twice more. 7. Add 0.1 volume of 3M NaOAc and 2.5 volumes of EtOH & incubate overnight at -20C. Collect the RNA by centrifugation at 7KRPM, 20 min., 4C (GSA or GS3 rotor), drain the pellet dry, and dissolve in 25ml STE- SDS. Precipitate by adding 3ml 3M NaOAc and 75ml EtOH, incubate at -70 for 1hr, the repellet 10KRPM, 4C, 20min., (SS34 or HB4 rotor), drain and dry the pelletes. 8. Dissolve the RNA in 10ml STE-SDS. Measure the absorbance of a 1:100 dilution of the sample & calculate the RNA concentration assuming that an A260 of 1 = 40ug/ml RNA. Pacelabs >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: EXONUCLEASE III FOOTPRINTING Exo III footprinting is a method for determining the site of binding for a protein on a DNA sequence. In order to use this procedure, you must have: purified protein, the DNA sequence of the cloned DNA in question, and filter-binding data indicating the general region of binding (or at least good reason to believe the binding site is within a relatively small region). Before starting the actual exo III footprinting, you need to isolate singley end-labeled probes of both strands containing the binding region, dissolved in ddH2O to 10,000 - 20,000 CPM/ul. These probes should be tested by DMS reactions (use 1/10 of the total amount of each probe - the rest of the DMS-treated probe is used for MW markers during the footprinting) for purity and to be sure the probes are the correct fragments. Labeling at 3' overhangs is OK, but changes in the digestion pattern may occur. 3' overhang second-cuts are NOT allowed - exo III will not reliably digest from the 3' overhang. Once you have these probes, you must determine the correct amount of exo III required for complete, but not over, digestion of EACH PROBE, empirically by titration. With this information you are ready to begin footprinting assays. Both the titration & footprinting are explained here. Exo III footprints (i.e.the boundary for the protein nearest to the labeled end) are seen as bands intermediate between the undigested probe and the complete digestion products. Unlike in the exo III titration, where the DNA steadily progresses from undigested to completely digested, with decreasing binding protein the DNA should pass directly from undigested to completely digested, since for each molecule of DNA, either the ends are covered by nonspecific binding or not. Any intermediate band(s) are either footprints or evidence that something in the protein sample is inhibiting the exo III. If the binding protein titration looks like the exo III titration, then it is probably due to inhibition of exo III activity by the protein sample. If only a few intermediate bands occur, they are most likely footprints, especially if they are strong signals. Exonuclease III - Boeringer Mannhiem - 160u/ul Store at -20C. TMK buffer 2X footprint buffer ------------- -------------- 50% glycerol 100mM KCl [KCl] may be adjusted 50mM Tris pH8 20mM MgCl2 depending on the 50mM KCl 40mM Tris pH8 optimum determined by 10mM MgCl2 0.2mM EDTA filter-binding. NTP mix STOP solution ----------- --------------- 3.3mM ATP 0.1M EDTA 3.3mM GTP 0.6M NH4OAc 3.3mM CTP 20ug/ml sssDNA Exonuclease III titration 1 - For each probe, mix (working on ice): 440ul ddH2O 50ul 5mg/ml BSA (nuclease-free) 500ul 2X footprinting buffer 50ul NTP mix 10ul probe DNA Divide the mix into 10 reactions of 100ul each, in eppendorf tubes. 3 - Do 8 1:2 serial dilutions of stock exo III (2ul TMK + 2ul exo III, mix, go to the next dilution). 4 - With all samples starting on ice, the experimental time course is : tube # 1 2 3 4 5 6 7 8 9 10 -------------------------------------------------------- --> 37C 0min 1 2 3 4 5 6 7 8 9 Exo III(1ul) 10 11 12 13 14 15 16 17 18 19 100ulSTOP->ice 20 21 22 23 24 25 26 27 28 29 -------------------------------------------------------- units exoIII 320 160 80 40 20 10 5 2 1 0 (2ul) 5 - When all the samples are done, add 100ul phenol:chloroform to each & vortex. Spin 1-2 min and transfer the upper phases to tubes containing 600ul EtOH. Freeze the samples in a dry-ice:EtOH bath for 5 min, then spin for 5 min. Discard the supernatants, and add 750ul ice-cold 70% EtOH to each tube, refreeze and spin as before. 6 - Resuspend each sample in 5ul sequencing dye and run a 6% sequencing gel. Run an appropriate distance for the size of the DNA used (broomophenol blue to bottom for 150bp, XC to bottom for 400bp). Dry & autoradiograph the gel (overnight exposure with screens at -70C). Choose the best digest for future use - you want digestion to the half- way point in the DNA, leaving 2 or 3 major bands near the middle of the molecules. Overdigestion results in degradation of the remaining, single-stranded, DNA. For 3' overhang labeled DNA, complete digestion of the labeled DNA strand may occur (rether than digestion to the half-way point) since exo III will not digest the unlabeled strand from the 3' overhang. In this case, use the lowest concentration of exo III that reliably gives total digestion of the labeled strand. Footprinting 1 - Working on ice, mix the same reaction mix as before for each probe. Divide the mix into 10 tubes each of 100ul. Prepare a dilution of Exo III if required. You should use the same dilution of exo III that gave optimum digestion in the titration. 2 - Prepare 1:2 dilutions (8 of them) of the binding protein to be tested in TMK. With all samples starting on ice, the time schedule is : tube # 1 2 3 4 5 6 7 8 9 10 --------------------------------------------------------- 1ulprotein->37C 0min 1 2 3 4 5 6 7 8 9 1ulexoIII 10 11 12 13 14 15 16 17 18 19 100ulSTOP->ice 20 21 22 23 24 25 26 27 28 29 --------------------------------------------------------- [protein] used 1X 1 1 1 1 1 1 1 1 0 - - - -- -- -- --- --- 2 4 8 16 32 64 128 256 --------------------------------------------------------- 3 - Phenol:chloroform extract, EtOH precipitate, 70% wash, dry, resuspend & run the 6% sequencing gel as before. Dry the gel and expose as indicated by experience from the exo III titration. Repeat the footprinting using ddH2O in place of NTP mix. JWB, unpublished Shalloway, et al 1980 Cell 20:411-422 >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: THE FIGURE-EIGHT TEST The figure-8 test is a test that identifies M13 clones that contain complementary sequences in their viral ssDNA. This occurs when the two clones contain the same region of DNA inserted in opposite orientations. This can be used to identify clones containing the same DNA fragment inserted in opposite directions, to identify clones containing nonidentical but overlapping complementary regions, or to find clones containing portions of DNA also found (complementary) in a larger clone. A good positive control for this test is M13mp8 against M13mp9, since they contain a 33 bp region inserted in opposite orientation (the polylinker). If properly done, this mixture will show a faint figure-8 band upon electrophoresis. NOTE: Broken circles are usually faintly visible in front of the circular ssDNA, and do not interfere with the test. NOTE: The viral DNA does not interfere because it is all (+) strand. 1. Prepare ssDNA from each strain to be tested by mixing 0.4ml stock virus (1-5 X 10/12 pfu/ml) with 50ul 2%SDS and 50ul 10X tracking dye. Vortex. 2. Mix the strains to be tested against each other (do this in duplicate) by adding 20ul of each of the two ssDNAs in a small microfuge tube and vortex. Add 100ul of sterile mineral oil to each tube, and centrifuge 5 sec. 3. Store one tube of each duplicate set at 4C. Incubate the other in a 68C water bath for 1 hr to allow the annealing of any complementary regions. 4. Run the duplicate samples side-by-side on a 1-1.5% agarose gel. Also run a lane containing M13mp8 ssDNA, as a standard. Those lanes containing a band with reduced mobility in the heated samples, but not in the unheated samples, are those with figure-8s'and therefore contain complementary sequences. Messing,J. 1983 'Recombinant DNA Techniques' Meth. in Enz. (in press) Messing,J. 1982 'New M13 Vectors for Cloning' NEN Booklet >>> From jwbrown@crab Wed Jan 15 02:45:08 Subject: FILTER BINDING ASSAY DNA fragments bound by RNA polymerase (or other DNA-binding proteins) can be retained on nitrocellulose filters, allowing the quantitation of binding activity of a specific DNA. This procedure is based on the fact that double stranded DNA will not bind nitrocellulose, but protein does. Complexes of DNA and RNA polymerase are formed then filtered through nitrocellulose - only DNA bound by the RNA polymerase will be held in the filter (since the enzyme is stuck), the remaining DNA is washed from the filter. Bound DNA is eluted by denaturing the enzyme, and analysed by gel electrophoresis. 1X FBB Elution buffer -------------- ---------------- 100mM KCl 0.2% SDS 10mM MgCl2 20mM Tris, pH8 0.1mM EDTA 0.1mM DTT NTP mix 10mM Tris, pH8 ------------------------- 50ug/ml BSA 3.33mM @ ATP, GTP, & CTP Phenol:chloroform Adjustment of [KCl] in 1X --------------- FBB may be required depending 50% TE sat'd phenol on the binding protein. 48% chloroform 2% isoamyl alcohol 1. Using a restriction map, decide on 4-8 restriction enzymes that give 5-12 bands of distinct MW for your DNA. Use these enzymes to cut your DNA, then end-label the DNA using any standard procedure (50-100uCi), then dissolve the final labeled DNA in 250ul ddH2O. For each labeled DNA: 2. Boil 5 (one is a spare) small (10-15mm) nitrocellulose filter circles for 10 minutes, and allow to cool while continuing with the experiment. 3. Prepare 4 eppendorf tubes with 2ul labeled DNA and 100ul 1X FBB. Add 0, 0.2, 1, and 5ul RNA polymerase to tubes 1-4 respectively, and incubate at 30C (or another temperature, depending on the RNA polymerase) for 5 minutes. 4. Add 5ul NTP mix, and continue the incubation for another 5 minutes, then transfer to ice. 5. GENTLY and SLOWLY filter each sample drop-wise through a boiled nitrocellulose filter, then wash the filter by filtering 500ul ice-cold 1X FBB. 6. Place the filter in a scintillation vial containing 400ul elution buffer and incubate 1 hour at 37C with shaking. 7. Collect the elute from the scintillation vial and add 50ul 2.5M NaOAc. To both the filtrate and eluate, add 400ul phenol:chloroform, vortex, spin, and collect the upper (aqueous) phase. To each, add 1ml EtOH, freeze 5 min in a dry-ice:EtOH bath, then spin 5 min. Discard the supernatant, and dry the pellet in a roto-vac. 8. Redissolve the pellets in 20ul each 2X tracking dye, and run on a 5% PAGE (or other gel, depending on the DNA fragment sizes). Dry the gel and autoradiograph overnight with screens (-70C). If desired, or required, scan the gel lanes with GELSCAN or another densitometer, and quantitate the intensities of each band. The results can then be analysed by hand (don't forget to correct for differences in labeling!) or by the FILTER program. Usually, eluates from complexes with the lowest amounts of RNA polymerase show the highest specificity and the lowest % retention. JWBrown, unpublished. Jones & Reznikoff 1977 J.Bact. 132:270-281 Hinkle & Chamberlin 1972 J.Mol.Biol. 70:157-185 >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Giant plasmid mini-prep For most purposes, such as sequencing, T7 transcription, subcloning, etc, the time & expense of CsCl:ethidium bromide purification of plasmid DNAs is not required. This procedure is essentially a scaled-up miniprep procedure that, because of the extensive organic extractions, is of sufficient purity for most purposes. If the DNA doesn't cut when you're done, or you need to get rid of the last of the RNA, you can purify this DNA using CsCl:EtBr, as usual (see CsCl purification of plasmid DNA). Sucrose buffer 25% SUCROSE (w/v), 50 mM Tris-HCl pH 8.0 LYSOZYME SOLUTION : 5 ml 416 ul 1M Tris-HCl (pH 8.0) 41.6 mg Lysozyme 1.6 ml 0.5 M EDTA (pH 8.0) 3 ml ddH2O TRITON LYSING SOLUTION : 2% Triton 10 mM Tris-HCl pH8.0 62.5 mM EDTA pH 8.0 1) Start a 10 ml culture of desired strain in a drugged medium and allow to grow for 4-6 hours (mid to late log phase). 2) From this inoculate 500 ml of drugged media with 2 ml of cells and allow to grow at 37¡C over night. 3) Transfer the culture to two 250 ml centrifuge bottles and cool on ice for 15 min. 4) Spin cells down in GSA rotor at 7.5K, 4¡C, for 10 min. 5) Pour off the supernatant, drain the bottles, and wipe the walls of the bottle with a Kimwipe. 6) Resuspend the cells (both pellets) in 5 ml Sucrose Buffer (the suspension will be almost opaque) and transfer to a 25ml 60Ti tube. 7) Add 5 ml of 10 mg/ml Lysozyme in TE Buffer and mix the tube gently. 8) Leave on ice for 15 min. 9) Add 10 ml of Triton lysing solution to the cells, seal the tube with Parafilm and mix by inverting the tube gently. 10) Leave on ice for 20 min. 11) Spin the tube at 35K, 30 minutes at 4C in the 60Ti rotor (be sure its precooled). 12) Pour the supernatant into a 35ml Corex tube and seal with Parafilm. 13) Heat the tube to 60¡C for 5 min. to ppt. the proteins. The solution should become cloudy. 14) Spin tube 10 min. 4¡C at 10K. 15) Transfer the supernatant to two 30 ml Corex tubes and extract each with 10 ml TE saturated phenol (pH 8.0). (The tubes should be sealed with a silicone #3 stopper and shaken for 30 sec. Since this is an organic extraction pressure may build up in the tube, so remember to burp the tube during the extraction and be careful when opening the tube. Spin the tubes in the table top clinical centrifuge for two min. to separate the two phases. Always wear gloves when working with phenol and/or chloroform.) 16) Repeat the extraction two times with 1:1, phenol:chloroform, and then with 10ml of chloroform. 17) Divide into two corex tubes & isopropanol ppt. the DNA with 4ml 7.5M NH4OAc and 28 ml isopropanol in each sample. Chill the tubes in ice for 15 min. 18) Pellet the DNA by centrifugation at 10K, 4¡C, for 20 min. 19) Resuspend the pellets in a total of 5 ml of 1X TE. 20) Transfer to a 15 ml screw cap dispo tube and add 50 ul of 10 mg/ml RNaseA. Seal the tube with its cap and secure the cap with Parafilm. 21) Incubate the tube for 1 hour at 37¡C. 22) Add 5 ml of saturated phenol to the tube and shake vigorously for 30 sec. and spin as above (15). 23) Transfer the aqueous phase to a 15 ml Corex tube and extract with 5 ml 1:1 , phenol : chloroform. Then extract with 5 ml of chloroform. 24) Isopropanol ppt. with 0.4 vol 7.5M NH40Ac and 2 vol. isopropanol. 25) Spin out the DNA and rinse the pellet with 70% ethanol. 26) Dry the tube walls with a Kimwipe and decicate the pellet lightly. 27) Resuspend the pellet in 1 ml 50 mM tris-HCl (pH 8.0). Quantitate the DNA by diluting 4 ul of your sample into 1 ml of H2O and reading the absorbance at 260 nm and 280 nm. Figure an A260 of 1.0 = 40 ug of DNA. PaceLabs Dirk Hunt, personal communication >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Genomic DNA isolation STE buffer pH 9.0 10mM Tris-HCl 100 mM NaCl 1 mM EDTA pH the solution to 9.0 with NaOH STE saturated Phenol 1) Melt the solid phenol in a 50 degree water bath. 2) Add 20% of the above STE Buffer (v/w). 3) Shake until the two become an emulsion. 4) Allow the phases to separate. 5) Decant off the aqueous phase. 6) Add Hydroxyquinolin to 0.1% (w/v) and Beta-mercaptoethanol (BME) to 0.2% (v/v). 7) Add the same amount of STE as you did in step 2) and store at 4 degrees. ( It may take more than one time of mixing to an emulsion to saturate the phenol) 50mM Tris-HCl, 0.1 mM EDTA 30% 4-Aminosalicylate,5% SDS (10 ml) 3 g 4-Aminosalicylate 5 ml 10% SDS 1) Grow 500 ml of cells to late log phase in appropriate medium (this will yield 2-3 g of E. coli in a rich medium). 2) Spin cells down in 250 ml centrifuge bottles(7.5K in GSA rotor; 20 min.) You will want a well-packed pellet. 3) Drain the pellet well and wipe the sides of the bottle with a Kimwipe. 4) Resuspend 1 gram of cells in 15 ml high pH STE in a screw cap bottle.(Many plastics are trashed with phenol and chloroform, Nalgene and polypropylene are OK, but polycarbonate is not.) 5) Add 2 ml 10 mg/ml lysozyme in H2O (make this fresh each time), incubate at 37¡C for 5 min. (the suspension should become less turbid). 6) Add 3ml 30% 4-aminosalicylate, 5% SDS; and gently mix for 1 min. 7) Hold at 70¡C for 10 min. The suspension should become completely clear. 8) Transfer to a 50 ml dispo centrifuge tube containing 20 ml phenol saturated with high pH STE buffer and put on the octopus for 30 min. at room temperature. (You may need to add more STE to maintain sample volume.) 9) Spin the tube in a clinical centrifuge at top speed for 10 min. 10) With a wide mouthed 25 ml pipette, remove the aqueous phase to a fresh tube containing 25 ml phenol. (Genomic DNA is very susceptible to shear damage.) 11) Repeat the extraction and phase separation 2 times. 12) Transfer the aqueous phase to a 40ml Nalgene Oak Ridge tube and extract with 20 ml of chloroform for 10 min. on the octopus. Spin 5 min. in clinical centrifuge. 13) Bring the aqueous to 20 ml with high pH STE and add 1 ml of RNase A* at 50 mg/ml (the RNase solution should be heated to 95¡C for 10 min. prior to use).*Use only disposable labware when working with RNase A 14) Transfer to a dialysis bag and dialize for 2-3 hours against 500 ml high pH STE buffer at room temperature. (Be sure to check bag for leaks prior to use.) This will allow the ribonucleotides to escape the bag. 15) Transfer to a dispo tube and add proteinase K to 50 ug/ml and SDS to 0.5%. ( 1.25 mg protenase and 1.25ml 10% SDS for 25 ml ). 16 ) Incubate for 1 hour at 50¡C. 17) Extract with 20 ml of high pH STE saturated phenol; followed by 20 ml of 1:1 phenol : chloroform ; and finally with 10 ml of chloroform. 18) Split the aqueuos phase into two corex tubes and add 1ml 3.0 M NaOAc, and 20 ml absolute ethanol to each tube. Chill in the -20 freezer for at least 1 hour. 19) Spool the DNA out of the tube with a Pasture pipette and transfer the DNA to an Oak Ridge tube containing 2-3 ml/(gram of original cells) of 50 mM Tris-HCl, 0.1mM EDTA (pH 8.0). Use the octopus at a slow setting or a slow reciprocal shaker to resuspend the DNA. Do not vortex. It may take several hours to a few days to resuspend the DNA. 20) Determine the concentration of the DNA by UV absorbance (A260 and A280). It is a good idea to try a restriction digest on the DNA and run it out on an agarose gel. Make sure that the DNA cuts well and that there is little RNA contamination. 21) Store at 4¡C. For long term storage, add a drop of CHCl3 to retard any bacterial and/or fungal growth. The chloroform can be evaporated from samples by a brief incubation at 37¡C. Pacelabs >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: GLYOXAL GELS RNA can be completely denatured by glyoxalating the bases in the presence of DMSO, thus prohibiting base pairing. RNAs denatured in this way will migrate in agarose or PAGE gels without interference because of secondary structure. Therefore, with this system, the molecular weights of RNAs can be closely estimated by comparing them to known standards (usually glyoxalated E.coli RNA) without the use of methylmercuric hydroxide. However, samples treated in this way should be relatively 'clean', so that the glyoxal is not depleted by contaminanting junk. Strict RNA-free technique should be used during this technique. All buffers, plasticware, and glassware should be autoclaved before use (except DMSO and glyoxal), and the electrophoresis comb and tray should be cleaned with EtOH. DON'T TOUCH THE TEETH ON THE COMB! If desired, 15mM Na iodoacetate can be incorporated into the gel. 0.2M NaH2PO4, pH 7.0 - 2.76 grams/100ml & pH with 5M NaOH 7M Glyoxal - Glyoxal is usually supplied at 7M. Before use, deionize by stirring with 10g mixed resin ion exchanger per 20ml, and store in small aliquots at -20C. 100X GE buffer (per liter) final conc. 69.0 grams NaH2PO4 0.5M 134 grams Na2HPO4-7H2O 0.5M - 1M PO4 total Adjust to pH 7.0 with 50% NaOH GE - diluted from 100X stock (10ml/l) Acridine orange stain (per 500ml) 5 mg acridine orange 10ug/ml 0.2922 grans NaCl 10mM Sample preparation: 1. Mix the following in a 500ul microfuge tube: 25ul DMSO (optional, if more RNA volume is needed) 2.5ul 0.2M NaH2PO4, pH 7.0 7.1ul 7M glyoxal 15.3ul RNA sample 50ul total volume 2. Incubate capped at 50C for 1 hr. 3. Add 5ul 10X tracking dye and load into the gel. Gel: 1. Use a 1-2.5% agarose gel (or 5-12% PAGE) made in GE. The gel should be poured while hot - about85C. 15mM Na iodoacetate can be added if RNase activity in the gel is a problem. 2. Recirculate the buffer by running the gel with the buffer trays on top of stir-motors, with gentle stirring, and with a peristaltic pump exchanging buffer between the trays. 3. Electrophorese at 1-5 V/cm until the dye reaches about 3/4 of the way down the gel. 4. Stain the gel for 1 hr in acridine orange, then destain by soaking in ddH2O for 1-1.5 hr. Examine under a UV light. To photograph, use Kodak Wratten filters; #12 for both green and red bands, #40 for green bands (DNA),and #29 for red bands (RNA). McMaster and Carmichael 1977 PNAS USA 74:4835 Maniatis, et al 1982 "Molecular Cloning" CSH Davis, et al 1980 "Advanced Bacterial Genetics" CSH pp156-158 >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Hybridization procedure #2 This procedure has been optimized for the use of oligonucleotide probes or short "fragmented" probes i.e. hydrolzed RNAs or nick translated small restriction fragments. 100X Denhardts (per 100ml) 2.0 gram BSA 2% 2.0 gram Ficoll 2% 2.0 gram PVP-360 2% 10% SDS - 10 grams per 100ml 20X SSC per liter 3M NaCl 175.3 grams 0.3M Na citrate 88.2 grams Adjust to pH 7 with 10N NaOH Hybridization fluid per 100ml 1X Denhardt's 1ml 100X 20mM NaH2PO4, pH7.6 2ml 1M 1mM DTT 200ul 500mM 5X SSC 25ml 20X 100ug/ml poly(A) 10ml 1mg/ml 10% dextran sulfate 20ml 50% (Add SDS to 0.1% when using an RNA probe.) Wash sol'n #1 - 2X SSC, 0.1% SDS Wash sol'n #2 - 1X SSC, 0.1% SDS Wash sol'n #3 - 0.1X SSC, 0.1% SDS 1. Put the filter to be hybridized to in a seal-a-meal bag, & add 5-10ml (10ml, unless the filter is small) of hybridization sol'n. Squeeze out all the bubbles & seal. Sometimes it helps to "roll" out the bubbles with a pipette. Be SURE that the bag is well sealed & won't leak! 2. Incubate at 65C for 1-2hr. This is best done in a water bath, & if the filter is vertically oriented the bubbles will float to the top, away from the filter. No shaking is required. 3. Cut off a corner of the bag & add the radio-labeled probe. If the probe is double-stranded DNA, preheat the probe in a boiling water bath for 2 min., & quick chill on ice before adding to the bag. Hybridize at 65C overnight. 4. Wash twice for 5 min. at RT in 40ml wash sol'n #1 at RT for 5min. each. This is done by cutting off a corner of the bag & draining all of the hybridization sol'n away (use a pipette to roll out the last of the sol'n) & then refilling the bag, via pipette or syringe, with the wash sol'n, & incubating. Check the filter with a geiger-counter; if the filter seems "cold", stop at this point & put the filter on film (see below, step 8). 5. Wash with 20ml Wash sol'n #2 at 65C for 15 min. 6. Wash with 20ml Wash sol'n #3 at 65C for 10 min. 7. Drain the last wash sol'n, & use a pipette to "roll-out" the last of the fluid, then reseal the bag without trapping air. 8. Autoradiograph as usual. Pace (Dirk) , personal communication Amersham Nytran specification sheet >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Preparing 5' end-labeled alkaline hydrolyzed RNA The best way to obtain high specific activity labeled rRNAs is to hydrolyze the RNA by alkaline treatment at high temperature, followed by 5' end-labeling with PNK & gamma-32P-ATP. This way, you get many 32P's incorporated into each rRNA molecule. This is also useful in heterologous probings, because the similarities of some of the RNA fragments will be much higher than the overall similarity for the heterologous rRNAs. After alkaline hydrolysis, phosphatase treatment is not needed because the fragments contain 5' hydroxyl's and 3' phosphates. REMEMBER TO USE RNase-FREE TECHNIQUE, ESPECIALLY AFTER THE HYDROLYSIS STEP. USE ONLY DEPC-TREATED H2O. 10X Kinase buffer TE-SDS ------------- ----------- 0.5M Tris,pH9.5 10mM Tris, pH8 0.1M MgCl2 1mM EDTA 50mM DTT 0.1% SDS 50% glycerol 1. Dilute 0.1-5 ug of RNA to 120ul in ddH2O, then divide into 4 30ul aliquots. 2. Add 30ul 100mM NaHCO3, pH9, to each sample and start the incubation at 95C. Remove one tube at 5 min. intervals (i.e. at 5, 10 15 and 20 min.) & transfer to ice. 3. Add 6ul 3M NaOAc and 180ul EtOH to each tube and pool the samples. Incubate in a dry ice:EtOH bath for 10min., centrifuge 10min., drain and dry the samples in a rotovac. 4. Redissolve the dry sample in: 5ul 10X PNK buffer 1ul (100uCi) gamma 32P-ATP 43ul ddH2O 1ul Polynucleotide kinase 5. Incubate at 37C for 30min., then add another 1ul of kinase & incubate 37C for 30min. 6. Add 10ul 100mM EDTA and 5ul of column dye. Separate the sample on a G50 column (in TE-SDS), collecting 20 15drop fractions. 7. Count 1ul of each fraction, & pool the highest fractions from the first peak. Store at -20C. Bryan James, personal communication >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: 5' End-labeling Alkaline Hydrolyzed rRNA The best way to obtain high specific activity labeled rRNAs is to hydrolyze the RNA by alkaline treatment at high temperature, followed by 5' end-labeling with PNK & gamma-32P-ATP. This way, you get many 32P's incorporated into each rRNA molecule. This is also useful in heterologous probings, because the similarities of some of the RNA fragments will be much higher than the overall similarity for the heterologous rRNAs. After alkaline hydrolysis, phosphatase treatment is not needed because the fragments contain 5' hydroxyl's and 3' phosphates. REMEMBER TO USE RNase-FREE TECHNIQUE, ESPECIALLY AFTER THE HYDROLYSIS STEP. USE ONLY DEPC-TREATED H2O. 100mM Tris, pH 9.5 Denaturation buffer per 1ml 10mM Tris, pH8 10ul 1M 0.1mM EDTA 1ul 100mM 0.19mg/ml spermidine-3HCl 7.5ul 100mM 10X PNK buffer 0.5M Tris,pH9.5 0.1M MgCl2 50mM DTT 50% glycerol 1. Dissolve 5-10ug of ribosomal RNA in 25ul ddH2O. Add 25ul 100mM Tris, pH9.5 and incubate at 95C for 3min. 2. Add 70ul denaturation sol'n, & heat for 3 min. at 50C. Chill on ice. 3. Working on ice, mix in: 148ul ddH2O 0.5ul (75uCi) gamma-32P-ATP 30ul 10X PNK buffer 10units (1-2ul) polynucleotide kinase 4. Incubate 37C 30min., then add 33ul 3M Na acetate and 660ul EtOH & incubate 5min. in a dry ice:EtOH bath. Centrifuge 5min., drain, and dry the RNA pellet. 5. Dissolve in 100ul ddH2O. Count 1ul to to determine incorporation. Store at -20C. Elizabeth S. Haas, personal communication Charles J. Daniels, personal communication >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: rRNA sequencing using reverse transcriptase Sequencing of rRNAs, primarily 16S rRNA at this time, is a useful method for quickly determining the phlyogenetic placement of an organism. Because of the presence of highly conserved regions in the 16S rRNA, universal oligonucleotide primers are available for sequencing portions of the 16S rRNAs from about any organism. These primers are situated in downstream of regions that are quite variable; these are sequences which are quite useful for phylogenetic analysis. The method described here is, of course, also aplicable for 23S rRNA sequencing provided appropriate oligonucleotide primers are used. The method can even be used for sequencing abundant mRNAs - in this case, it might be useful to modify the method by using end-labeled primers rather than incorporating radiolabeled dNTPs into the reaction mixes. The most important part of this procedure is the RNA isolation. Use RNase-free technique! Otherwise, the procedure is not essentially different from chain termination sequencing from DNA templates. If the sequences are difficult to interprete, try re-precipitating the RNA in 2M NaCl (0C overnight) to remove low MW RNAs. Also try decreasing the primer concentration in the sequencing reactions 5-fold (i.e. use an 0.02mg/ml stock concentration). 5X hybridization buffer 5X RT buffer 100mM KCl 250mM Tris, pH8.5 250mM Tris, pH8.5 50mM DTT 50mM MgCl2 RT dilution buffer Sample buffer 50mM Tris, pH8.3 86% formamide 2mM DTT 10mM EDTA 50% glycerol 0.08% xylene cyanol 0.08% bromophenol blue Gel fix - 10% methanol, 10% glacial acetic acid, 2% glycerol Oligonucleotide primers - dissolved in ddH2O to 0.1mg/ml primer 519 5'-GWAATACCGCGGCKGCTG-3' primer 915 5'-GCCCCCGYCAATTCCT-3' primer 1391 5'-GACGGGCGGTGTGTRCA-3' PREPARATION OF NUCLEOTIDE MIXES 1. For dNTPs, dissolve 10mg in 1ml 10mM Tris, pH8.3. For ddNTPs, dissolve 4umoles in 300ul 10mM Tris, pH8.3 2. Dilute 4ul of this into 2ml 10mM Tris, pH 7.4, and read the absorbance at that (d)dNTPs absorption maximum - the (d)dNTP concentration can be calculated as: (absorbance)(dilution factor) ----------------------------- = concentration in mM mM extintion coefficient The constants needed are: nucleotide absorption max (nm) mM extinction coeff dATP 259 15.2 dCTP 272 9.1 dGTP 253 13.7 dTTP 267 9.6 dITP 249 12.2 [S]dATP 259 14.8 ddATP 259 15.2 ddCTP 272 9.1 ddGTP 253 13.7 ddTTP 267 9.6 3. Dilute the initial solution to 10mM by adding 10mM Tris, pH8.3. 4. To the alpha-thio-dATP solution, add 1/1000 volume (1ul/ml) 1M DTT. 5. Store at -70C. 6. Make small (~500ul) working stocks of each of the following in 10mM Tris, pH8.3: 1mM ddCTP, 1mM ddGTP, 1mM ddTTP, and 0.1mM ddATP. 7. Six nucleotide mixes are needed for RT sequencing (numbers are in ul): G-mix C-mix T-mix A-mix no-dd mix chase ------------------------------------------------------------ 10mM dGTP 10ul 10 10 10 10 140 10mM dCTP 10 10 10 10 10 140 10mM dTTP 10 10 10 10 10 140 10mM [S]dATP 5 5 5 5 5 - 10mM dATP - - - - - 140 1mM ddGTP 7.6 - - - - - 1mM ddCTP - 12 - - - - 1mM ddTTP - - 12 - - - 0.1mM ddATP - - - 5 - - 10mM Tris,pH8.3 353 349 349 356 361 833 1M MgCl2 - - - - - 7 100mM DTT 4 4 4 4 4 - ------------------------------------------------------------ 8. Divide the mixes into 20ul aliquots, except chase mix, which gets 70ul aliquots, and store at -20C. RNA PREPARATION RNA can be prepared by standard methods, depending on the cell type. High molecular weight RNA should be removed from total RNA preparations by NaCl precipitation, or rRNA should be isolated by phenol extraction & ethanol precipitation from ribosomal pellets (i.e. S100 pellets). rRNA should be dissolved in ddH2O at a concentration of about 2mg/ml. REVERSE TRANSCRIPTASE SEQUENCING REACTIONS The sequencing reactions are easiest done in sets of three primers (i.e. 3 sets od sequencing reactions). This allows you to use the standard three universal primers quickly. 1. Prepare the following tubes: Hybrid mixes A B C ----------------------------------------------------------- 5X hybridization buffer 1.4ul 1.4ul 1.4ul template RNA (2mg/ml) 4ul 4ul 4ul primer A (0.1mg/ml) 1.5ul - - primer B (0.1mg/ml) - 1.5ul - primer C (0.1mg/ml) - - 1.5ul ----------------------------------------------------------- 2. Incubate at 65C for 2 min., then slow-cool to room temperature over about 10min., then prepare the following tubes, on ice: Pre-mixes A B C ----------------------------------------------------------- [alpha-35S]dATP (30uCi) dry dry dry 5X RT buffer 6ul 6ul 6ul Hybrid mix A 6ul - - Hybrid mix B - 6ul - Hybrid mix C - - 6ul 1000units/ml RT 6ul 6ul 6ul ----------------------------------------------------------- The RT is diluted to 1u/ul (1000u/ml) in RT dil'n buffer just prior to addition. 3. Still working on ice, prepare the following mixes for each premix (in is best to have the dNTP/ddNTP mixes pre-aliquoted into tubes, ready for the addition of the pre-mixes): Sequencing reactions Reaction A B C ------------------------------------------------------ A pre-mix 3ul 3ul 3ul A-mix 2ul 2ul 2ul G pre-mix 3ul 3ul 3ul G-mix 2ul 2ul 2ul T pre-mix 3ul 3ul 3ul T-mix 2ul 2ul 2ul C pre-mix 3ul 3ul 3ul C-mix 2ul 2ul 2ul Control pre-mix 3ul 3ul 3ul no dd mix 2ul 2ul 2ul ------------------------------------------------------ 4. Incubate at room temperature for 5 min., then 30 min. at 55C. 5. Add 1ul chase mix (10:1 chase mix:1000u/ml RT) & incubate 15 min., 55C. 6. Dry the samples in a rotovac, and redissolve each sample in 10ul sample buffer. Electrophorese on 6% sequencing gels: two loadings are best, with the second started as the first samples xylene cyanol reaches 2/3rds of the way to the bottom of the gel. Run the gel until the bromophenol blue of the seconf loading reaches the bottom of the gel. Soak the gel in Gel soak for 15 min., then dry and autoradiograph. Try a 2 day exposure. PaceLabs protocol sheet (Steve Giovanonni) Lane 1985 PNAS 82:6955 Sanger 1977 PNAS 74:5463 Biggin 1983 PNAS 80:3963 >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: ISOLATING M13 CLONES AND PREPARING INFECTED CULTURES These are some of the basic techniques used in harvesting clones, preparing stock virus preparations, and preparing infected cultures from M13 and M13 derivatives. The infected cultures can be used for a variety of purposes, such as ssDNA isolation, figure 8 tests, rapid RF isolation, etc. YT can be substituted with LB, L broth, or B broth. For subsequent preparations, 100ul stock virus can be substituted in place of a picked plaque. For large-scale RF isolations, this technique can be scaled-up to several liters without modification, if care is taken to assure good aeration. Preparing infected cultures: 1. Inoculate 20ml YT (in a 125ml flask) with 0.4ml of an overnight culture of E.coli JM103. Incubate at 37C with vigorous agitation for 1- 1.5 hr. 2. Using a sterile pasteur pipette, remove a plug of agar containing a well isolated plaque. 3. Transfer the plaque to the 20ml culture, and rinse off the inner walls of the pipette tip by drawing the culture back-and-forth through the pipette. 4. Incubate as before for 5-7 hr. This is then an infected culture, which can be stored for several days at 4C until continuing. 5. Fill a 1.5ml microfuge tube with infected culture, and spin for 1 min. 6. Transfer the supernatant to a sterile 1.5ml microfuge tube, and heat for 15 min. at 65C. This is the stock virus preparation, and should have a titer of about 1-2 X 10/12. 7. Store at 4C. Rapid ssDNA isolation: 1. Mix 45ul stock virus with 5ul 2% SDS in a 500ul microfuge tube. 2. Add 5ul 10X tracking dye and separate on a 1-1.5% agarose gel. Rapid RF DNA isolation: RF DNA can be isolated from the pellet in step 6 by any rapid plasmid isolation technique, such as the boiling method described in the CSH Molecular Cloning manual (pages 366-367). Zinder and Boeke 1982 Gene 19:1-10 Messing,J. 'Recombinant DNA Techniques', Meth. in Enz. (in press) Messing,J. 'New M13 Vectors for Cloning' NEN booklet 1982 >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Molecular Cloning Manual (Maniatis) The Molecular Cloning lab manual by Maniatis, et al (CSH 1982) has proven to be a very valuable guide to basic recombinant DNA techniquiques. A variety of methods from the manual have been tried in our lab, usually with success. Below are some of these techniques which have been found to work well with little or no modification MAINTENANCE OF BACTERIAL STRAINS Use 100ug/ml ampicillin in plates. RESTRICTION ENZYMES - i.e. the buffers RAPID END-LABELING OF DNA (pp115-116 AGAROSE GEL ELECTROPHORESIS (pp150-162) Use 0.25ug/ml ethidium bromide when incorporated into the gel and buffer. Also, we use trays constructed from plexiglass. MINIGELS (pp163) You can also run minigels on microscope slides using 5-6 ml agarose. DNA PAGE (pp173-177) GLYOXAL GEL ELECTROPHORESIS OF RNA (pp200-201) For Northerns, treat blots with 20mM Tris, pH8, at 100C for 10 min after baking to reverse the glyoxylation. FORMALDEHYDE GELS FOR RNA (pp202-203) STAINING NORTHERNS WITH METHYLENE BLUSE (pp206) not particularly sensitive SOUTHERN TRANSFER (pp383-386) TRANSFORMATION USING CALCIUM CHLORIDE (pp250-251) RAPID PLASMID ISOLATION (boiling method) (pp366-367) At step #9, it is easier to incubate 30 min. at -70. We've had poor luck trying to cut or sequence this DNA in some strains of E.coli (i.e.JM103). PLASMID AMPLIFICATION ON RICH MEDIA APPENDICES Preparation of organic reagents (pp438-439) Liquid media (pp440-441) Media containing agar or agarose Antibiotics Preparation of stock solutions Solutions RNase that is free of DNase (pp451) Commonly used electrophoresis buffers (pp454) Commonly used gel loading buffers (pp455) Xylene cyanol is optional. Preparation of dialysis tubing (pp456) Drying down 32P-dNTPs (pp457) Purification of nucleic acids (pp458-460) Concentration of nucleic acids (pp461-463) DNA has sometimes been lost during butanol concentration. Chromatography through sephadex G50 (pp464-465) Spun column procedure (pp456-467) Use a good centrifuge to minimize speed variation Quantitation of DNA or RNA (pp468-469) Autoradiography (pp470-472) We usually use a single intensifying screen. NOTES: * Notice the handy concentration chart for acids and bases on pp445. * It is very handy for all of the people in the lab to be using solutions prepared to the concentrations described in this manual, so that borrowing back-and-forth is easier. >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Culturing Methanogens Methanogenic bacteria are extreme obligate anaerobes and require H2 and CO2 for growth. Care should be taken at all times to prevent the introduction of air or oxygenated solutions into the cultures. Also, since the cultures are grown under positive pressure, be careful to prevent embarrassing mistakes (i.e. vent before removing the crimped top, and HOLD ON to the plunger whenever using a syringe). NOTES * Stationary M. vannielii has an A600 of 0.8-0.9. M. fervidus grows to a lower density, & the methanobacteriales grow to a higher density. * In minimal media (ER), M. vannielii (& most other methanogens)has a doubling time of about 4.8 hr. Vitamin solution (per liter) 2 mg biotin 2 mg folic acid 10 mg pyridoxine HCl 5 mg thiamine HCl 5 mg riboflavin 5 mg nicotinic acid 5 mg DL-Ca pantothenate 0.1 mg vitamin B12 5 mg PABA 5 mg lipoic acid FILTER STERILIZE! Mineral I (per liter) liter 10X 6 grams K2HPO4 60 grams Mineral II (per liter) liter 10X 6 grams KH2PO4 60 grams 6 grams (NH4)2SO4 60 grams 12 grams NaCl 120 grams 2.6 grams MgSO4-7H2O 26 grams 0.16 grams CaCl2 1.6 grams Trace minerals (per liter) liter 10X 24 mg Na2SeO3 0.24 grams 0.1 grams CoCl2 1 gram 24 mg Na2MoO4-2H2O 0.24 grams 24 mg NiCl2-6H2O 0.24 grams 10ml con HCl 100 ml (CAUTION! SODIUM SELENITE IS EXTREMELY TOXIC!) Reducing agent (per 400ml) 1. Boil and cool 400ml dH2O. 2. Add 5g cysteine-HCl. 3. Adjust pH to 12 with NaOH pellets. 4. Add 5g Na2S-9H2O. 5. Divide into 150ml serum bottles, filling them about half full. 6. Purge with N2 and stopper. 7. Autoclave 15 min. Resazurin - 2 pellets per 88ml ddH2O Prepare 1 liter ER medium as follows: 1. Dissolve the following in a 2 liter flask: 880ml ddH2O 2.7g NH4Cl 5.0g NaHCO3 1.25g Na2CO3 2.5g Na acetate 2ml fresh 0.5% FeSO4 (0.5g/100ml) 50ml mineral I 50ml mineral II 10ml trace minerals 10ml vitamins 10ml reazurin 2. Heat to a light boil over a burner. While the medium is heating, purge 4 @ 1 liter bottles at 15 PSI using bent needles and a 40:60 ratio of CO2:H2 (2:1 on the meter). 100ml vaccine bottles can also be used. 3. Pour the media into the bottles while still very hot (250ml @ per 1 liter bottle, 20ml @ per 100ml bottle) and continue purging for a few minutes. 4. Plug the bottles without letting any air in, and crimp on aluminum tops. Autoclave 25 min. in a pan of water. 5. After the media has cooled to RT, add 1ml reductant per 100ml media (2.5ml per 250ml, 0.2ml per 20ml). 6. Pressurize the bottles to 15 PSI (40 PSI for 100ml bottles) with a 40:60 CO2:H2 gas mix (2:1 on the meter), using a sterile needle and gas sterilizing syringe. 7. When the medium becomes colorless, inoculate with 2ml of a stationary phase (or log phase) culture and incubate at the optimal growth temperature for 2 or 3 days. Pressurize twice daily as before. Lenny Hook, personal communication Hook, et al "Modified Anaerobe Chamber" unpublished >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: FERMENTING METHANOGENS with the New Brunswick Microferm 24 liter FIRE HAZARD!!!!! POTENTIALLY EXPLOSIVE!!!! TOXIC GAS! POISON! DEFINITELY have someone show you how to do this! Directions for growing fermenter batches in the OSU Microbiology Fermentation Facility fermenters. See the directions for preparing methanogen medium for the recipes for ER. Vitamin solution (per liter) 2 mg biotin 2 mg folic acid 10 mg pyridoxine HCl 5 mg thiamine HCl 5 mg riboflavin 5 mg nicotinic acid 5 mg DL-Ca pantothenate 0.1 mg vitamin B12 5 mg PABA 5 mg lipoic acid FILTER STERILIZE! Mineral I (per liter) liter 10X 6 grams K2HPO4 60 grams Mineral II (per liter) liter 10X 6 grams KH2PO4 60 grams 6 grams (NH4)2SO4 60 grams 12 grams NaCl 120 grams 2.6 grams MgSO4-7H2O 26 grams 0.16 grams CaCl2 1.6 grams Trace minerals (per liter) liter 10X 24 mg Na2SeO3 0.24 grams 0.1 grams CoCl2 1 gram 24 mg Na2MoO4-2H2O 0.24 grams 24 mg NiCl2-6H2O 0.24 grams 10ml con HCl 100 ml (CAUTION! SODIUM SELENITE IS EXTREMELY TOXIC!) Reducing agent (per 400ml) 1. Boil and cool 400ml dH2O. 2. Add 5g cysteine-HCl. 3. Adjust pH to 12 with NaOH pellets. 4. Add 5g Na2S-9H2O. 5. Divide into 150ml serum bottles, filling them about half full. 6. Purge with N2 and stopper. 7. Autoclave 15 min. Resazurin - 2 pellets per 88ml ddH2O GAS SYSTEM PREPARATION 1 - Connect the appropriate gas cylinders (usually CO2 and H2), with regulators, to the copper gas tubes that run to the gas mixer. With valve E off (pointed down), turn the gas cylinders on and adjust the regulator pressure to 25 PSI. Check for leakage (i.e. listen for a hissing sound, or if H2 is being used, use the gas leak detecter). Turn the gas cylinders off. 2 - Turn valve A (behind the fermentor/air incinerator) toward the anaerobic gas line (i.e. to the left), and close valve Z (the aerobic gas line). Open valve Y (anaerobic gas exhaust) and turn valve C (next to the gas mixer) to the right. 3 - On the back of the white fermentor (Fermentation Design 30l fermentor) in the SE corner of the lab, close valves B and D, to insure there is no exchange of gas between the fermentors. 4 - Check the valves by following the open gas lines from the gas cylinders to the vertical anaerobic gas exhaust. You should be able to trace the correct gas route just by following the open valves, and all lines connecting to the lines should be closed off by valves. Make sure the metal airstone is screwed onto the main sparger at the bottom of the inside of the fermentor, and check that the bolts holding the fermentor to the lid are tight. 5 - Open the main sparger valve and the exhaust valve (all other valves on top of the fermentor should be closed). 6 - Turn on the gas cylinders. Gas should flow in the lines, and the balls in the gas mixer should float. Adjust the gas mixing to 15 CO2 and 8 H2 (for CO2:N2, use 10 and 12 respectively). 7 - Turn off the gas cylinders, valve E, the main sparger, and the exhaust. Medium preparation 1 - To the dry, clean fermentor, add the following through the large screw-capped opening at the top of the ferm lid using a powder funnel: 54 grams NH4Cl 100 grams NaHCO3 25 grams Na2CO3 50 grams Na acetate 40 mls 0.5% FeSO4 100 mls 10X mineral I 100 mls 10X mineral II 20 mls 10X trace minerals 200 mls vitamin sol'n 80 mls resazurin sol'n (or 2 tablets) 2 - Add ddH2O to 20 liters, and close the fermentor. Stir for a few minutes at 200 RPM to dissolve the ingredients. Alternately, the dry ingresients can be added to the fermentor vessel before it is bolted to its lid, eliminating spillage & mess. Then, the liquids can be poured through the opening as usual. 3 - Turn on the gas cylinders and open valve E (turn to the left), the main sparger, exhaust (in that sequence). Allow to bubble and stir for at least 15 minutes, to purge the medium and head space of oxygen. Leave the agitation on. 4 - Close exhaust, main sparger, valve E (point down), and valve Y (in sequence). Check once again that valve Y is CLOSED. (Leaving valve Y open during the sterilize cycle will fill the outlet lines with water from condensed stream & flood the ceiling when the gas line is opened!) 5 - Open the top & bottom main steam lines (in corner of room), steam line, and middle water line (both are in back of fermentor, next to the floor). Check the inlet and exhaust filter jacket steam lines (on either side inside the fermentor) - they should both be on at all times. Switch the "STERILIZATION/OFF/TEMP CONTROL" switch to "STERILIZE", then turn ON all the switches in the front of the fermentor labeled "STERILIZE", and turn OFF the seal steam condenser water valve, and open the sampling steam (on the front). Be sure the temperature setting is all the way up (clockwise). Check all of the valves on the top of the fermentor - they must all be closed. 6 - When the temperature reaches 110C, one at a time open the valves for the harvest, main sparger, exhaust, and inoculation ports for 5 minutes each. CAREFULLY watch the medium level in each case - if the level drops, CLOSE THE PORT & go the next one. When the temperature approaches 121C, begin timing the sterization cycle (20-30 min). 7 - Turn off all the STERILIZE switches and the sample steam valve, turn temperature control on and set to the required incubation temperature, open the seal stream water condenser, and turn cooling control to manual. 8 - When the temperature drops to 100C, open valve E, the main sparger, exhaust, and valve Y (in sequence), then turn on the exhaust condenser switch (just above the exhaust valve). Continue stirring at 200 RPM. This drives out any oxygen in the head space of the hot medium. 9 - When the temperature reaches about 10C above the incubation temperature, turn the cooling switch to automatic. 10 - Connect a preautoclaved piece of plastic tubing with a vaccine top on it to the inoculation port. Swab the vaccine top with 70% EtOH and insert a needle barely into the top. Open the inoculation valve and close the exhaust valve, allowing the pressure to vent through the needle to drive the air from the tubing. 11 - Remove the needle and open the exhaust port. Using a double- ended needle and pre-pumped up bottles, inject 200ml reducing agent into the inoculation port. Be sure the liquid drains into the fermentor, rather than just sitting in the tubing, then close the inoculation port valve. If any substrate but CO2:H2 is to be used, such as MeOH or methylamine, inject it at this point. 12 - Turn off the exhaust, main sparger, valve E, gas cylinders (in sequence). 13 - The medium is ready to use after it turns dark grey (a few hours). INOCULATION AND FERMENTATION 1 - Prepare 1 liter (3 X 300ml) of dense culture in 1 liter Wheaton bottles. Pump the bottles up to 15 PSI. 2 - Turn on (in sequence) the gas cylinders, valve E, the main sparger, exhaust, and agitation (100 RPM for bugs grown without shaking, 300 RPM for bugs grown with shaking). 3 - Swab the bottle stoppers and inoculation vaccine top, open the inoculation port, and inject the inoculum into the port. Close the inoculation port. 4 - 20ml of a sterile 15% Na2S solution should be injected in the same way through the inoculation port twice daily. 5 - The growth of the bugs can be monitored through the window, or by removing small samples through the sample port. If the sample port is used, be sure to turn on the sampling steam for 5 minutes to resterilize the port. HARVESTING METHOD 1 - BATCH CENTRIFUGATION 1 - Connect a N2 gas cylinder to the anaerobic collection carboy using tygon tubing, and purge the carboy for about 15-20 minutes at low pressure. Be sure the other 2 inlets into the carboy are open. Leave the N2 gas on. 2 - Connect the carboy to the harvest port with tygon tubing. Close the exhaust port and open the harvesting port valve. As pressure builds up in the fermentor, the culture will be forced into the carboy. 3 - When the carboy is full, close the harvest port and open the exhaust port. Clamp off the tubing from the fermentor to the carboy, and then simultaineously turn off the N2 gas cylinder and clamp off the carboy exhaust, then clamp off the carboy N2 gas inlet tubing, thus sealing off the carboy. 4 - Turn off the gas cylinders, valve E, main sparger, exhaust, valve Y, top main steam lines, and the water inlet line (at the back of the fermentor on the floor - turning off the water at the knob by the door to the room also turns the water to the ice machine off). Turn the power switch off. 5 - Move the carboy and N2 gas cylinder down to room 468, next to the left side of anaerobe chamber side A. Connect the harvesting tube to the rearmost inlet port on the side of the anaerobe chamber. Reconnect the N2 cylinder to the carboy, and turn the regulator pressure ALL THE WAY DOWN (VERY IMPORTANT!), and turn the gas cylinder on. Open the clamp on the N2-carboy tubing, and on the carboy-chamber tubing. 6 - Hold the inlet tubing inside the chamber over a steel centrifuge bottle (250ml), and open the valve on the inlet port. CAREFULLY increase the regulator pressure until a slow, even flow into the centrifuge bottles is obtained. 7 - When all the centrifuge bottles are full, close the inlet valve and clamp off the inlet tubing inside the chamber. Clamp off the carboy N2 inlet, thus again sealing off the carboy. Leave the regulator pressure on - with the carboy sealed off, no dangerous pressure can build up. 8 - Carefully and firmly screw on the centrifuge bottle lids and remove them from the chamber. Centrifuge at 5000 RPM for 10 minutes at 4C, then ship them back into the chamber. 9 - Decant the supernatants into a 4 liter waste bottle, and refill the bottles as before. Repeat until the entire culture is pelleted. While the bottles are centrifuging, syphon the waste ER out of the chamber using the other chamber inlet port. Be sure the port valves are closed when not in use. 10 - Resuspend the pellets in about 100ml waste ER, and wash the bottles out with another 100ml waste ER. Pool the cells in one of the bottles, seal the lid, and centrifuge as before. 11 - Ship the bottle into the chamber, and discard the supernatant. Using a spatula, transfer the pellet into a preweighed bottle that can be stoppered tightly. Add 5ml ER to the remaining pellet material, resuspend, and pool with the pellet. 12 - Stopper the bottle and screw on a lid. Remove the bottle from the chamber and weigh it. The yeild is 5 grams less that the net weight (since there's 5g of ER in the bottle). Store the cell paste at -70C. METHOD 2 - FILTRATION (for Ms.barkeri) 1 - Connect vacuum tubing from the sink aspirator/vacuum to the chamber outlet port. Inside the chamber, connect the same port to a 1 liter vacuum flash with a Buchner funnel. 2 - Connect the harveting tube from the carboy through the chamber inlet port. Inside the chamber, connect some tubing to the inlet port & clamp it over the funnel so that liquid will flow from the tube into the funnel. Connect the N2 cylinder to the carboy exhaust as described above. 3 - Place a piece of Whatmann #1 paper in the funnel, & wet the paper. 4 - Start the vacuum, & open the outlet port valve, allowing the vacuum to draw through the filter. 5 - Start the flow of medium into the funnel by opening the inlet port, unclamping the carboy harvest line & exhaust line, & BARELY opening the regulator, allowing very slight gas flow into the carboy,, pumping medium into the chamber. 6 - As the filtrate proceeds, adjust the medium flow rate to match the filtration rate. Do not be concerned about filling the vacuum flask - when it is filled, the excess fluid will be drawn through the tubing into the sink drain. Be sure the filtrate remains clear. 7 - If the filtration rate slows too much, stop the filtration, change the filter, & continue. 8 - Scrape the cells from the filter paper into a sealable glass bottle & store at -70C. METHOD 3 - CONTINUOUS FLOW CENTRIFUGATION See the material on the use of the continuous flow rotor for the sorval SS-34. Make the following modifications : 1 - The housing MUST be filled with oil - this is not optional. 2 - After the rotor has attained the set speed (10K or higher), VERY GENTLY flush the rotor with N2 for 5 min. 3 - Clamp off the rotor tubing, & reconnect the N2 gas to the carboy exhaust, then connect the carboy to the rotor inlet tubing. 4 - Gently pump the medium into the rotor using gas pressure - DO NOT RELY ON CENTRIFUGAL PUMPING - CONTROL THE FLOW RATE WITH A PINCH CLAMP ON THE ROTOR INLET TUBE. The flow rate should be ralatively fast, but the supernatant should be clear. If the sup is turbid, slow the pumping rate, & perhaps increase the rotor RPM (up to 15K). 5 - Be sure the culture is cold before starting. 6 - When finished, stop the rotor & remove the individual centrifuge tubes. Close the inlet & outlet of each by connecting the two with thin rubber tubing. Pump them into the chamber, discard the sups, & respuspend the pellets in 20ml waste ER. Transfer the slurry to an SS- 34 tube & pellet the cells 10K 10min 4C, & transfer them to an airtight glass bottle for storage at -70C. 7 - Take the rotor COMPLETELY apart, rinse & clean. This is particularly important since the sulfide in ER is corrosive. Lenny Hook, personal communication JWBrown, personal communication >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Micro-protein assay This micro-protein assay is not as accurate as other methods for assaying protein concentration, but it has the distinct advantage of requiring only trace amounts (less than 1ug) of your valuable protein. In many cases the accuracy (within a factor of 2) of the assay is more than sufficient. Another advantage of the assay is its relative ease and quickness. Amido black stain 0.25% Amido black 0.625g 45% MeOH 112.5ml 45% ddH2O 112.5ml 10% glacial HOAc 25ml Amido black destain same as the stain without the dye Cellulose acetate dissolving solution 80ml formc acid 10ml glacial HOAc 1ml 100% TCA Starting with 5mg/ml BSA (in saline), make a series of 1:2 dilutions in saline (100ul BSA + 100ul saline). 8 - 10 dilutions is fine. In duplicate, spot 1ul of each BSA dilution, and the samples to be tested to a strip of cellulose acetate. Allow the spots to air-dry. If the samples contain alot of sucrose or glycerol. Re-wet the spot with 2ul aliquots of ddH2O & allow to dry again. Repeat this 3 - 5 times. Stain in amido black stain for 10 minutes, then destain in several transfers of destain until the background is nearly white. Examine the intensities of the black spots. Compare the spots from your test protein with those of the standards to obtain an estimate of your protein concentration. If a more accurate estimate is required: Carefully punch out one of each duplicate spot & dissolve in 400ul dissolving solution Measure the A523 of each sample. Plot the BSA standards as a standard curve & interpolate the concentration of the test samples using this curve. Michael Thomm, personal communication JWBrown, unpublished data >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Precipitation of high MW RNAs by NaCl Often, it is desirable to obtain high MW RNA, free of degradation fragments of small RNAs (i.e. tRNAs), for either rRNA sequence analysis or Northern analysis of mRNAs. The method can also be used in reverse - small RNAs can be isolated free of high MW rRNA components. This can be accomplished by taking advantage of the differences in soluability of RNAs of different size in high concentrations of NaCl. High MW RNAs are insoluable, at 0C, in 2M NaCl, whereas low MW RNAs are soluable. Remember to use RNase-free technique! 1. Dissolve RNA to about 5mg/ml in ddH2O or TE, then add an equal volume of 4M NaCl. 2. Incubate overnight at 0C (i.e. in an ice bath at 4C). 3. Centrifuge for 10min. in an eppendorf microfuge at 4C, discard the pellet, and dissolve the high MW RNA in the original volume of TE. If you want the low MW RNA, dilute the supernatant in 9 volumes of 300mM NaOAC, then add 3X the new volume of EtOH. Incubate at -70 for 2hr, and centrifuge 15KRPM, 4C, 20min. (SS34 rotor), drain & dry the pellet and dissolve in a small volume of TE. 4. Measure A260 to determine the exact concentration of RNA (it should be ~80% in high MW RNA and ~20% in low MW RNA). Pace, personal communication >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: NICK TRANSLATION dsDNA can be in vitro radiolabeled with 32P by treatment with DNase I, DNA polymerase I, dNTPs, and a32P-dCTP. Several commercial kits are available for nick translation, but some laboratories prefer to nick translate from scratch to improve incorporation and save on costs. Of course, strict precautions must be used, as whenever mCi quantities of 32P is used. DNase I storage buffer (per 20ml) filter sterilize 50mM Tris, pH 7.5 - 1ml 1M stock 1mM DTT - 200ul 0.1M stock 10mM MgSO4 - 200ul 1M stock 50% glyerol - 10ml DNase I dilution buffer (per 20ml) filter sterilize 50mM Tris, pH 7.5 - 1ml 1M stock 10mM MgSO4 - 200ul 1M stock 1mM DTT - 200mM 0.1M stock 10X NT buffer per 10ml 0.5M Tris,pH 7.5 5ml 1M stock 0.1M MgSO4 1ml 1M stock 10mM DTT 1ml 0.1M stock 0.5mg/ml BSA 5mg BSA (nuclease-free) dNTP mix - 0.2mM @ dTTP, dCTP & dGTP, each from 2mM stocks. For labeling with alpha-32P dNTPs besides dATP, use the other 3 dNTPs in the dNTP mix. Stop sol'n per 20ml 20mM Na3 EDTA - 4ml 0.1M stock 2 mg/ml sssDNA - 4ml 10mg/ml stock (see hybridization Rx for sssDNA recipe) TE per liter 10mM Tris, pH 7.5 - 10ml 1M stock 1mM EDTA - 10ml 0.1M stock Column dye per 20ml 0.25% bromophenol blue - 50 mg 50% glycerol - 10ml 1/2 X TE - 10ml 1X Preparation of DNase I 1. Add 0.5ul 1mg/ml stock DNase I to 100ul dilution buffer on ice. 2. Add 0.5ul diluted DNase I to 100ul dilution buffer, again on ice. Final dilution is 1:40,000. Nick translation 1. Mix the following ingredients on ice: 2.5ul 10X NT buffer 2.5ul dNTP mix 50uCi alpha-32P-dATP 0.5ul diluted DNase I X ul DNA (1ug) Y ul ddH2O (to 20ul) 2. Add 0.1ul 2mg/ml DNA polymerase I. Mix and incubate at 14C for 3 hrs. 3. Add 25ul Stop sol'n. Column chromatography 1. Add 50ul TE and 10ul column dye to the sample. 2. Apply the sample onto a 0.7 X 20 cm G50 sephadex column in TE. 3. Run the column with TE, collecting the first 20 fractions, 12 drops each. Wash the column in TE until the blue dye is all washed out of the column. 4. Spot 2ul of each fraction onto 4-5mm squares of 3MM paper. Count each square in 4ml scintillation fluid. 5. Pool the G50 excluded peak (i.e. the first peak), which usually is at fractions 7-10. Store at -20C until use. CSH 'Molecular cloning' 1983 CSH molecular cloning workshop, attended by P. Hamilton Paul Hamilton, personal communication >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: Northern gels RNA can be completely denatured by glyoxalating the bases in the presence of DMSO, thus prohibiting base pairing. RNAs denatured in this way will migrate in agarose or PAGE gels without interference because of secondary structure. Therefore, with this system, the molecular weights of RNAs can be closely estimated by comparing them to known standards (usually glyoxalated E.coli RNA) without the use of methylmercuric hydroxide. In fact, since glyxylated DNAs and RNAs with the same molecular weights migrate the same, linear DNA, such as restriction fragments, can be used as MW standards for estimating the MWs of unknown RNAs. However, samples treated in this way should be relatively 'clean', so that the glyoxal is not depleted by contaminating junk. Strict RNase-free technique should be used during this technique. All buffers, plasticware, and glassware should be autoclaved before use (except DMSO and glyoxal), and the electrophoresis comb and tray should be cleaned with EtOH. DON'T TOUCH THE TEETH ON THE COMB! 0.1M NaH2PO4, pH 7.0 (10X GE) - 13.8 grams/liter Adjust to pH 7.0 with 50% NaOH 6M Glyoxal - Glyoxal is usually supplied at 6M (40% aqueous). Before use, deionize by stirring with 10g mixed resin ion exchanger (AG 501-X8) per 20ml, and store in small aliquots at -20C. Acridine orange stain (per 500ml) 5 mg acridine orange 10ug/ml 0.2922 grans NaCl 10mM 5X GE tracking dye (per 20ml) 2ml 10X GE 10mM PO4 10ml glycerol 50% 0.08 grams bromophenol blue 0.4% 8mlddH2O Sample preparation: 1. Mix the following in a 500ul microfuge tube: 2.7 ul 7M glyoxal 8.0 ul DMSO 1.6 ul 0.2M NaH2PO4, pH 7.0 5 ul or less RNA sample 2. Incubate capped at 56C for 1 hr. While the samples are incubating, prepare the gel & apparatus as described below. Gel: 1. Use a 1.0-2.0% agarose gel (or 5-12% PAGE) made in GE. The gel should be poured while hot - about 85C. 2. Recirculate the buffer by running the gel with the buffer trays on top of stir-motors, with gentle stirring, and with a peristaltic pump exchanging buffer between the trays. 3. Electrophorese at 1-5 V/cm until the dye reaches about 3/4 of the way down the gel. 4. If the gel isn't going to be transferred to nitrocellulose, stain the gel for 1 hr in acridine orange, then destain by soaking in ddH2O for 1- 1.5 hr. Examine under a UV light. To photograph, use Kodak Wratten filters; #12 for both green and red bands, #40 for green bands (native DNA, rRNA),and #29 for red bands (denatured RNA & DNA, non-rRNA). Northern Blotting glyoxylated RNA or DNA RNA normally has a low affinity for nitrocellulose since native RNA has extensive secondary structuring. Thus, older methods were developed for Northern blotting using DBM-activated paper to bind native RNA. However, glyoxylated nucleis acids are thoroughly denatured and will readily bind nitrocellulose. Therefore both RNA and DNA can be easily transferred from agarose gels to nitrocellulose, after glyoxylation. 1. Wet a pre-cut sheet of nitrocellulose in ddH2O, then soak in 20X SSC for at least 5 min. 2. Lay a wetted (in 20X SSC) 3MM paper wick over an upside-down gel tray in a dish of 20X SSC. Ovr this lay a piece of 3MM paper (again, pre- wetted in 20X SSC) cut exactly the same size as the gel. Roll a pipette over the paper to remove ALL of the air bubbles. 3. Place the gel directly after electrophoresis (i.e. untreated & unstained) face-down onto the wet 3MM paper. Again, roll out any trapped air bubbles with a pipette. 4. Place the nitrocellulose filter on top of the gel, being careful not to trap any air under the nitrocellulose (or roll them out as before). 5. Cover the filter with 2 layers of pre-wetted (20X SSC) 3MM paper cut the same size as the gel. For the last time, remove trapped air by rolling over the paper with a pipette. 6. Surround the gel on all four sides with strips or saran-wrap, so that the dish and wick are covered. The sara-wrap should butt up against the gel on all four sides. 7. Lay a thick stack of paper towels onto the top 3MM paper sheet. Over this, lay a glass plate large enough to cover the area of the gel. On top, place a 500ml flask filled with water, as a weight. 8. Replace the paper towels as they become wet, whenever convenient. The transfer is complete after 16-24 hrs. 9. Remove the nitrocellulose filter & allow it to air-dry. DO NOT WASH BEFORE BAKING! Bake at 80C in a vacuum oven for 2hr. 10. Bring 500ml 10mM Tris, pH8, to a boil over a bunsen burner, then pour this hot solution into a plastic tray. Drop the filter into the hot Tris, submerge the filter, then wait for the solution to cool. Remove the filter, air dry, and store wrapped in foil at RT until use. This step reverses any residual glyoxylation, allowing more efficient hybridization. Staining Northerns with Methylene blue After autoradiography, Northerns may be stained with methylene blue to visualize the transferred RNA. This is especially useful, since glyoxylated RNAs should not be stained before transfer to nitrocellulose from agarose gels. 1. Soak the nitrocellulose filter in 5% acetic acid for 15 min. 2. Transfer the filter to methylene blue stain (0.5M Na acetate, pH 5.2 + 0.04% methylene blue), and soak for 10 min. 3. Rinse the filter for at least 10 min. in ddH2O. McMaster and Carmichael 1977 PNAS USA 74:4835 Maniatis, et al 1982 "Molecular Cloning" CSH Davis, et al 1980 "Advanced Bacterial Genetics" CSH pp156-158 Thomas 1980 PNAS USA 77(9):5201 >>> From jwbrown@crab Wed Jan 15 02:45:08 1992 Subject: NITROCELLULOSE FILTER HYBRIDIZATION Filter hybridization is a method used to qualitatively determine the presence of homologous or complementary sequences in DNA or RNA, by allowing a denatured radio-labeled 'probe' nucleic acid (in solution) to anneal to the denatured nucleic acid (immobilized to a nitrocellulose filter) to be tested. The filters containing the immolbilized nucleic acid can be Southern blots, Northern blots, lifted colonies or plaques, or spot filters. For Northern blots or RNA probes, use strict RNase-free technique. For DNA-DNA hybridizations, poly(A) and blocking DNA may not be needed. At low stringency, blocking DNA may interfere with proper hybridization. Choose a blocking DNA that is most dissimilar to both DNAs being hybridized (i.e. herring sperm DNA for hybridizing procaryotic DNAs, or E. coli DNA for hybridizing yeast against soybean). Stringency can be adjusted from high (68C) to low (down to about 50C) by lowering the temperature. Hybridizations at high stringency, as described here, with DNAs of perfect homology, will require about 100 KCPM of 32P-probe DNA. Hybridizations at low stringency generally require more probe CPM. 20X SET (per liter) final conc. 175.3 grams NaCl 3.0M Adjust to 72.6 grams Tris-OH 0.6M pH 8.0 14.9 grams Na2EDTA 40mM with HCL 50X Denhardts (per 100ml) 1.0 gram BSA 1% 1.0 gram Ficoll 1% 1.0 gram PVP-360 1% 20% SDS - 10 grams per 50ml 10% Na pyrophosphate - 5 grams per 50ml (boil to dissolve) 10 mg/ml poly(A) - 10 mg plus 1.0ml ddH2O (store -20C) 50ug/ml sonicated herring sperm DNA - 5 mg plus 5ml ddH2O, stored at - 20C and sonicated (Micro-tip, 3-5 min. at max frequency) in 1ml aliquots before use. Sol'n A (per 200ml) 40ml 20X SET 4X 160ml ddH2O Sol'n B (per 100ml) 20ml 20X SET 4X 20ml 50X Denhardts 10X 0.5ml 20% SDS 0.1% 59.5ml ddH2O Sol'n C (per 50ml) 10ml 20X SET 4X 10ml 50X Denhardts 10X 0.25ml 20% SDS 0.1% 50ul 10mg/ml poly(A) 10ug/ml 0.5ml 10% Na pyrophosphate 0.1% 28.7ml ddH2O Sol'n D (per 100ml) 20ml 20X SET 4X 20ml 50X Denhardts 10X 0.5ml 20% SDS 0.1% 1.0ml 10% Na pyrophosphate 0.1% 100ul 10mg/ml poly(A) 10ug/ml 58.5ml ddH2O Sol'n E (per 300ml) 45ml 20X SET 3X 1.5ml 20% SDS 0.1% 3.0ml 10% Na pyrophosphate 0.1% 250.5ml ddH2O Sol'n F (per 500ml) 25ml 20X SET 1X 2.5ml 20% SDS 0.1% 5ml 10% Na pyrophosphate 0.1% 468ml ddH2O Sol'n G (per 200ml) 10ml 10X SET 1X 190ml ddH2O Handle the filters with gloves! 1. Wet the filter by floating, then sinking it, in 200ml sol'n A at room temperature. Let the filter soak for 30 min. 2. Transfer the filter to