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{\rtf1\mac\ansicpg10000\uc1 \deff12\deflang1033\deflangfe1033{\upr{\fonttbl{\f0\fnil\fcharset256\fprq2{\*\panose 02020603050405020304}Times New Roman;}{\f4\fnil\fcharset256\fprq2{\*\panose 02000500000000000000}Times;} {\f5\fnil\fcharset256\fprq2{\*\panose 02000500000000000000}Helvetica;}{\f7\fnil\fcharset256\fprq2{\*\panose 020b0503030404040204}Geneva;}{\f12\fnil\fcharset256\fprq2{\*\panose 02020502060305060204}New York;} {\f26\fnil\fcharset256\fprq2{\*\panose 020b0806080604040204}Chicago;}}{\*\ud{\fonttbl{\f0\fnil\fcharset256\fprq2{\*\panose 02020603050405020304}Times New Roman;}{\f4\fnil\fcharset256\fprq2{\*\panose 02000500000000000000}Times;} {\f5\fnil\fcharset256\fprq2{\*\panose 02000500000000000000}Helvetica;}{\f7\fnil\fcharset256\fprq2{\*\panose 020b0503030404040204}Geneva;}{\f12\fnil\fcharset256\fprq2{\*\panose 02020502060305060204}New York;} {\f26\fnil\fcharset256\fprq2{\*\panose 020b0806080604040204}Chicago;}}}}{\colortbl;\red0\green0\blue0;\red0\green0\blue255;\red0\green255\blue255;\red0\green255\blue0;\red255\green0\blue255;\red255\green0\blue0;\red255\green255\blue0; \red255\green255\blue255;\red0\green0\blue128;\red0\green128\blue128;\red0\green128\blue0;\red128\green0\blue128;\red128\green0\blue0;\red128\green128\blue0;\red128\green128\blue128;\red192\green192\blue192;}{\stylesheet{\sl240\slmult0 \widctlpar\adjustright \f12\cf1\cgrid \snext0 Normal;}{\*\cs10 \additive Default Paragraph Font;}{\s15\sl240\slmult0\widctlpar\tqc\tx4320\tqr\tx8640\adjustright \f12\cf1\cgrid \sbasedon0 \snext15 footer;}{\s16\sl240\slmult0\widctlpar \tqc\tx4320\tqr\tx8640\adjustright \f12\cf1\cgrid \sbasedon0 \snext16 header;}}{\info{\author Travis Glenn}{\operator Travis Glenn}{\creatim\yr2000\mo10\dy31\hr14\min21}{\revtim\yr2000\mo10\dy31\hr14\min22}{\version2}{\edmins12}{\nofpages19} {\nofwords4281}{\nofchars24405}{\*\company SREL}{\nofcharsws29971}{\vern99}}\margr1440 \facingp\widowctrl\ftnbj\aenddoc\hyphhotz0\aftnnar\hyphcaps0\viewkind1\viewscale109 \fet0\sectd \linex0\titlepg\sectdefaultcl {\headerl \pard\plain \s16\sl240\slmult0 \widctlpar\tqc\tx4320\tqr\tx8640\adjustright \f12\cf1\cgrid {\f5 Appendix D\tab \tab Getting Old DNA}{ \par }}{\headerr \pard\plain \s16\sl240\slmult0\widctlpar\tqc\tx4320\tqr\tx8640\adjustright \f12\cf1\cgrid {\f5 Appendix D\tab \tab Getting Old DNA}{ \par }}{\footerl \pard\plain \s15\sl240\slmult0\widctlpar\tqc\tx4320\tqr\tx8640\adjustright \f12\cf1\cgrid {\field{\*\fldinst {PAGE }}{\fldrslt {\lang1024 12}}}{\f5 \tab \par }}{\footerr \pard\plain \s15\sl240\slmult0\widctlpar\tqc\tx4320\tqr\tx8640\adjustright \f12\cf1\cgrid {\field{\*\fldinst {PAGE }}{\fldrslt {\lang1024 11}}}{\f5 \tab \par }}{\headerf \pard\plain \s16\sl240\slmult0\widctlpar\tqc\tx4320\tqr\tx8640\adjustright \f12\cf1\cgrid {\f5 Appendix D\tab \tab Getting Old DNA}{ \par }}{\footerf \pard\plain \s15\sl240\slmult0\widctlpar\tqc\tx4320\tqr\tx8640\adjustright \f12\cf1\cgrid {\field{\*\fldinst {PAGE }}{\fldrslt {\lang1024 1}}}{\tab \par }}{\*\pnseclvl1\pnucrm\pnstart1\pnindent720\pnhang{\pntxta .}}{\*\pnseclvl2\pnucltr\pnstart1\pnindent720\pnhang{\pntxta .}}{\*\pnseclvl3\pndec\pnstart1\pnindent720\pnhang{\pntxta .}}{\*\pnseclvl4\pnlcltr\pnstart1\pnindent720\pnhang{\pntxta )}} {\*\pnseclvl5\pndec\pnstart1\pnindent720\pnhang{\pntxtb (}{\pntxta )}}{\*\pnseclvl6\pnlcltr\pnstart1\pnindent720\pnhang{\pntxtb (}{\pntxta )}}{\*\pnseclvl7\pnlcrm\pnstart1\pnindent720\pnhang{\pntxtb (}{\pntxta )}}{\*\pnseclvl8 \pnlcltr\pnstart1\pnindent720\pnhang{\pntxtb (}{\pntxta )}}{\*\pnseclvl9\pnlcrm\pnstart1\pnindent720\pnhang{\pntxtb (}{\pntxta )}}\pard\plain \qc\sl240\slmult0\widctlpar\adjustright \f12\cf1\cgrid {\b\f7 Getting DNA from Old Dead Stuff}{\f7 \par }\pard \sl-360\slmult0\widctlpar\adjustright {\b\f7 \par }\pard \qc\sl-360\slmult0\widctlpar\adjustright {\b\f7 Travis Glenn}{\f7 \par }\pard \qc\sl240\slmult0\widctlpar\adjustright {\f7 \par }\pard \sl240\slmult0\widctlpar\tx4680\adjustright {\f7 Savannah River Ecology Lab\tab Department of Biological Sciences \par Drawer E\tab University of South Carolina \par Aiken,SC 29802\tab Columbia, SC 29208 \par e-mail Glenn@srel.edu\tab e-mail: Travis.Glenn@sc.edu \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par \par }\pard \sl-360\slmult0\widctlpar\adjustright {\b\f7 \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \tab There are two majo r questions regarding getting DNA out of museum specimens: 1) what's the best tissue to sample, and 2) what's the best way to get DNA out of that tissue? For dried bird skins, the short answers are: 1) the easiest well-dried tissue to get, and 2) the e asiest extraction methods you are comfortable doing. \par \par \tab Most of my efforts have centered on extracting DNA from small bits of bird skin. I have also extracted DNA from bone, cartilage, muscle, & feathers. I have used a variety of "extraction" techniques: chelex, phenol-chloroform, silica, and commercial kits. Each technique has some advantages as well as disadvantages, and I provide a few comments on those in this document. Below I present the protocols I have used in my attempts to isolate DNA from old dead stuff. I also present some comments and information that may prove useful if you wish to really understand what I have done, or are foolish enough to attempt this type of research yourself. \par \par \tab I have now obtained a couple hundred samples from museum b ird skins. Although I think the protocols I present are good protocols, I am not convinced that they are yet optimal. If one is given sufficient time and free reign to obtain DNA from museum birds, I would recommend rehydrating the skin, everting a wing , and collecting bone, muscle, and cartilage. I would then isolate the DNA using QiaAmp Tissue kits or good old proteinase K digestions followed by PCI extractions and microconcentration. The rehydration and eversion of skins is tricky & you would be wel l advised to seek the advice of someone with considerable experience in handling museum bird skins (not me). I would seek advice from Kevin Winker (now at the Univesity of Alaska). QiaAmp kits are nice because they come with most solutions prepared (less ening chances of introducing contaminants from your lab) & have few steps. \par \par \tab I have shown for museum Whooping Cranes that there is significant deterioration of DNA over time. Most of my PCR efforts have focused on mitochondrial DNA (mtDNA). I have also at tempted amplification of microsatellites, and those results are largely congruent with the mtDNA results. Among the specimens>80 years old, no amplicons were derived from about one third of the specimens, amplicons of only about 50 bp were obtained from another third, and amplicons of 50 - 164 bp were obtained from the final third. Amplicons of>200 bp were obtained from very few specimens>80 years old. \par \par \tab In general, the tissues sampled and protocols used for DNA isolation will probably not greatly af fect the results obtained. However, attention to minute details of the protocols chosen may greatly affect the level and detection of contamination. Thus, I am providing the explicit protocols I used to obtain DNA from museum specimens, and detect conta mination. \par \par \tab Finally, I should probably warn you that my general philosophy is to regard every step in a protocol as a potential point for introducing contamination and that everything possible should be done to minimize and detect contamination. I could not disagree more strongly with the statement of Thomas et al. (1990:101): "This study demonstrates the accuracy and }{\b\f7 routine nature}{\f7 of the use of museum specimens ..." (emphasis mine). Routine contamination control measures are usually inadequate and may lead to conflicting results (see footnote 5 of Thomas et al. 1990). Working on "ancient", or more appropriately for me, "antique" DNA deserves a healthy fear of contamination. This fear can manifest itself as bold belief of unlikely results (e.g., foot note 5 of Thomas et al. 1990; Woodward et al. 199}{\f7\cf6 3}{\f7 ) or pure paranoia (see below). You will be the judge of my manifestation. \par \par }\pard \sl-360\slmult0\widctlpar\adjustright {\f7 Travis Glenn November 1996 (address update 9/98), formerly at}{\b\f7 \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 Laboratory of Molecular Systematics; MRC 534, MSC; Smithsonian Institution; Washington, DC 20560. e-mail: Glenn@onyx.si.edu \par }{\b\f7 \page Overview }{\f7 \par \par \tab The process obtaining DNA from museum specimens can be outlined with the following steps: 1) preparing an overall strategy, 2) preparing to sample, 3) collecting samples, 4) prepar ing to extract DNA, and 5) extracting DNA. Sections 1, 2, and 4 are mostly advice for anyone considering such projects. The remaining sections detail protocols that I have used. \par \par }{\b\f7 I. An Overall Strategy}{\f7 \par \par }{\f7\ul \tab Project design}{\f7 - In addition to the normal considerations (e.g. sampling design), an ideal project would have the following characteristics: 1) the PCR primers are already designed and tested, or can be designed and tested away from the samples, 2) there is no way for DNA contamination from the researchers to be confused with DNA obtained from the study specimens, 3) large numbers of samples are available, 4) independent facilities for sample collection, preparation, and amplification are available, and 5) results can be validated, ideally, with independent collaborators who can replicate results. \par \par }{\f7\ul \tab Pre and Post PCR Laboratories}{\f7 - The importance of physically separating pre- and post-PCR manipulations can not be stressed enough. PCR amplicons are easily transmitted. It is difficult to slow the flow of small amounts of previous PCR amplicons from making their way into subsequent reactions. The most effective method is to physically separate pre- and post-PCR manipulations. Increasing the separation of the manipulation s increases contamination control. \par \tab Ideally, samples are taken, DNA is extracted, and PCR reaction components are assembled in one facililty (or more). Then, thermocycling and analysis are completed in another separate facility. Reagents and material sh ould be kept completely separately for the facilities. All reagents and materials that must be shared, should move one way, from the first lab to the second (cf. Cano 1993). Unfortunately, if the same people are involved in both facilities, true one-way movement is impossible. Thus, one must consider the timing of primer design and optimization. \par \par }{\f7\ul \tab PCR primer design and optimization}{\f7 - Ideally, PCR primers can be designed and tested at a facility that is completely isolated from the facility where the old DNA will be handled. If that is not possible, then as few individuals as possible should be used for primer design and testing. Additionally, it is best to temporally isolate primer design and testing (presumably using modern samples) from museum sample collection, DNA extraction, and DNA amplification. \par \par }{\f7\ul \tab dUTP and UDG}{\f7 - The incorporation of dUTP into amplicons and the subsequent use of UDG is the most powerful method available to destroy previous amplicons, and thus control carry-over contamination (see L ongo et al. 1990, and Glenn and Braun 1992, for background information and protocols). Be forewarned, however, that U-DNA amplicons do not necessarily behave "normally" (e.g., see Glenn et al. 1994). The most profound effect that is not yet in the liter ature involves the apparent inefficiency of }{\i\f7 Taq}{\f7 DNA polymerase to incorporate dUTP relative to TTP. I have only noticed this effect for amplicons longer than 300 bp. My initial results indicate that this inefficiency can be overcome by simply increasing the incubation time at 72}{\f4 \u730\'fb}{\f7 C (1.5 - 2x normal has done the trick so far). \par \par }{\f7\ul \tab Think, rehurse, do}{\f7 - Care should be taken at every step to reduce the potential of contamination. Prepare and manipulate all negative controls as close to the samples as possible. Always handle samples, then negative controls. Give the negatives every possible chance to get contaminated. You want them the negatives to represent the "worst case", not the "best case". \par \par \par }{\b\f7 II. Preparing to Sample}{\f7 \par \par \tab All tissues should be collected with disposable single-use scalpels, syringes, and plasticware whenever possible. Reagents, tools, and stocks of disposable supplies used for museum specimens should be purchased new especially for such projects and should kept in laboratories free of potent ially contaminating DNA or samples until use. Reagents and supplies should not be obtained from general stocks of supplies (even in pre-PCR labs). \par \tab Re-usable tools (e.g., forceps, drill bits, etc.) should be soaked in or wiped down with>10% bleach between samples to destroy any DNA on the tools (Prince and Andrus 1992). Instruments and labware that can be baked, should also be baked at>300}{ \f4 \u730\'fb}{\f7 C for>2 hr. Exposing lab surfaces, when not in use, to ultraviolet light is also advisable. \par \par \tab Before sampling, you will want to compile a list of samples you want, a sampling data sheet, and order "fresh" sampling supplies. \par \par \par }{\b\f7 III. Collecting Samples}{\f7 \par \par }{\b\f7 \tab A. Sampling Bird Skins \par }{\f7\ul \par Overview}{\f7 \par \tab The basic gist is to collect a chunk of skin in the least destructive manner possible. I usually accomplished this by separating the feathers along the ventral suture and taking a skin sample that contained few associated feathers. I also opportunistica l ly took skin samples from other regions (often under damaged wings). Curators were usually present to witness the destructive sampling of the first specimen or three. After they had a feel for what I wanted & were reasonably sure of what I would take, I usually worked by myself. If there were any "problematic" specimens, I would always double-check with the curators to be sure that what I wanted to sample was ok with them. \par \tab Skins that still contained high levels of grease were very soft & yellowish-brown to dark brown. These skins did not seem to contain much DNA. "Good" skins were: dry, hard, and off-white. \par \par Equipment & Materials: \par }\pard \li360\sl240\slmult0\widctlpar\adjustright {\f7 Sample sheets \par Disposable scalpels \par Small (50 mL) beakers \par Forceps \par dH}{\f7\fs20\sub 2}{\f7 0 \par Bleach \par Kimwipes \par Disposable powderless gloves \par Ziplock Baggies \par Sharpies \par Optional: A big roll of clean paper (white or brown) \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par }{\f7\ul Procedure}{\f7 \par }\pard \fi-440\li440\sl240\slmult0\widctlpar\adjustright {\f7 \par 1) Place 2+ forceps into beaker with bleach diluted 1:1 with dH}{\f7\fs20\sub 2}{\f7 0. Fill another beaker with dH}{\f7\fs20\sub 2}{\f7 0. \par \par 2) Clean a surface to work on, wiping down with 10% bleach. If possible, tear off a piece of clean paper bigger than your bird specimens & work on this. \par \par 3) Get out a specimen (AKA old dead bird). \par \par 4) Record specimen number & data on record sheet. \par \par 5) Write specimen number on ziplock baggie. \par \par 6) Place one set of forceps into beaker of dH}{\f7\fs20\sub 2}{\f7 O. \par \par 7) Put on gloves & determine the best place to obtain skin sample. \par \par 8) Clean forceps with kimwipe & set aside. \par \par 9) Open new disposable scapel & break-off blade guard. \par \par 10) Grasp the desired piece of skin with the forceps and cut the sample - about 1 cm x 2}{\f7\super +}{\f7 cm. Take as much as is reasonable, including attached feathers, without damaging the specimen. \par \par 11) Place the skin sample in the ziplock. \par \par 12) Place the second set of forceps into the dH}{\f7\fs20\sub 2}{\f7 0. \par \par 13) Wipe-off the first set of forceps (the ones you just used) with a kimwipe and then place them into a beaker with diluted bleach. \par \par 14) Rewrap the scapel inside its original foil (for disposal or later use with modern samples). \par \par 15) Return the specimen to the appropriate drawer. \par \par 16) Repeat steps 3 - 15 for all remaining specimens. The surface should be wiped with a clean paper towel. However, you don't need to wipe the surface with bleach each time because you are }{\i\f7 not}{\f7 going to drop the skin sample. \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par \par }{\b\f7 \tab B. Drilling Bird Bones}{\f7 \par \par }{\f7\ul Overview}{\f7 \par \tab The basic gist is to collect at least 0.1 g of bone in the least destructive manner possible. Although, Dremel sells a variety of bits, I found the plain old drill bit to give the biggest (and therefore the best) sized bone "dust". Other bits designed for grinding turned the bone into a fine powder that was difficult to capture anywhere other than in my lungs. I also found that I used the lower speed (7,500 rpm) almost exclusively. The faster speed caused the bone to burn (which stinks & is unlikely to be good for the DNA). \par \tab Bones that were covered with brownish soft tissue which still contained high levels of grease were very soft & yellowish brown. I highly doubt that this bone will contain much DNA. "Good" bones were: dry, hard, off-white, and yielded dust that was quite white. \par \par Equipment & Materials: \par }\pard \li360\sl240\slmult0\widctlpar\adjustright {\f7 Sample sheets \par Dremel Cordless Handdrill Model 770 \par 1/8th inch drill bit \par Emery cutting wheels \par Aluminum Foil (cheap, non-heavy duty, is fine) \par 1.5 mL centrifuge tubes \par Disposable scalpels \par Small (50 mL) beakers \par dH}{\f7\fs20\sub 2}{\f7 0 \par Bleach \par Kimwipes \par Disposable powderless gloves \par Ziplock Baggies \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par }{\f7\ul Procedure}{\f7 \par }\pard \fi-440\li440\sl240\slmult0\widctlpar\adjustright {\f7 \par 1) Place forceps & drill bit into beaker with bleach diluted 1:1 with dH}{\f7\fs20\sub 2}{\f7 0. \par \par 2) Tear off a piece of aluminum foil at least 10 cm x 15 cm. \par \par 3) Get out a specimen (AKA old dead bird). \par \par 4) Record specimen number & data on record sheet. \par \par 5) Write specimen number on 1.5 mL tube. \par \par 6) Place forceps and drill bit into beaker of dH}{\f7\fs20\sub 2}{\f7 O. \par \par 7) Put on gloves & determine the best place to obtain bone sample. \par \par 8) Place aluminum foil under the location to be drilled/cut. \par \par If drilling: \par \par 9) Use forceps to retrieve drill bit from the bottom of the beaker of dH2O. Dry off the drill bit using a clean Kimwipe & secure the bit into the drill. \par \par 10) Dry off the forceps using a Kimwipe & place them onto another clean Kimwipe. \par \par 11) Using the sl ow speed & gentle pressure, drill into the bone. Drill until you reach the center of the bone (i.e. the open space in the middle). Note: the drill bit may get stuck occasionally. When this happens, try wiggling it gently. If the bit is still stuck, t urn off the drill & wiggle more aggressively, until it becomes free. \par \par 12) Use the forceps to "tap-off" bone dust that is still on the bone onto the aluminum foil. \par \par 13) Gently remove the aluminum foil from under the bone. \par \par 14) Make a crease & pour the bone dust into the labeled 1.5 mL tube (using the forceps to brush the bone dust, if necessary). I generally collect 200 - 500 "\u181\'b5 L" of bone dust. If you need more, then replace the aluminum foil under the bone & drill another hole or two. \par \par If collecting a "hunk": \par \par 9) Place a new emery wheel on the appropriate drill bit. Secure this into the drill. \par \par 10) Dry off the forceps using a Kimwipe & place them onto a clean Kimwipe. \par \par 11) Using the slow speed & gentle pressure, cut into the bone. Cut until you pass through the first layer of the bone (i.e. into the open space in the middle). Continue the cut by moving the drill around the bone (i.e. continue to cut from the outside in, not down and through the bone). Caution: the wheel may catch (i.e. get stuck) occasionally. When this happens, it can cause the drill to lurch out of your hands. Hold on tight! \par \par 12) Use the forceps to "tap-off" bone dust that is still on the bone onto the aluminum foil. \par \par 13) Gently remove the aluminum foil from under the bone. \par \par 14) Use the forceps to place the bone chunk into a labeled ziplock baggie. \par \par 15) Make a crease & pour the bone dust into the labeled 1.5 mL tube (using the forceps to brush the bone dust, if necessary). \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par \par }{\b\f7 \tab C. Sampling Bird Feet}{\f7 \par \par }{\f7\ul Overview}{\f7 \par \tab The gist is to collect a chunk of foot pad in the least destructive manner possible. DNA may be better preserved in the foot pad because this tissue is highly cornified (see Sawyer 199}{\f7\cf6 x}{\f7 ; N. Mundy pers. commun.). The problem with collecting foot pad tissue is that becau se it is so tough, when one liberates a chunk of it, one tends to send it flying across the room. Thus, the major challenge is to keep the foot pad clean and under control. I have had the best luck using a technique similar to the one outlined for skin s amples. However, I tend to put a piece of aluminum foil under & in front of the foot. That way, if the chunk of foot pad shoots off, it goes into the aluminum foil. I also attempting using some "bone clipers" (look like wire cutters). This worked ok, but the tissue chunks really flew. \par }\pard \fi-440\li440\sl240\slmult0\widctlpar\adjustright {\f7 \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par }{\b\f7 IV. Preparing to Extract DNA \par }{\f7 \par \tab Order "fresh" chemicals and supplies. When possible, order pre-made stocks (e.g., 1 M Tris pH 8, 0.5 M EDTA, 20% SDS, Proteinase K). Pour aliquots of your stocks into 50 mL disposable tu bes for use. These precautions will reduce the risk of you contaminating these solutions considerably. It is also advisable to make larger than expected quantities of your working solutions. Then, test a small aliquot of your solutions in a PCR negativ e control to ensure that everything is copasetic before you launch into extractions. \par \tab It may not be a bad idea to do a small sub-set of extractions before you do a huge number of sample extractions. However, if you do PCRs, it is best to use a locus diff erent from the one you want to screen. What if you don't have another locus? That's the conundrum, you can leap in head-first without looking (i.e. do no test extractions); or put your toe in the water (i.e. do some test extractions) & blow your plan to keep PCR products away from your samples until all the extractions are done. \par \par \par }{\b\f7 V. Extracting DNA \par }{\f7 \par \tab Although there are a large number of variations on the same basic themes, most DNA extraction procedures can be put into one of four categories: 1) denatur ation and dilution (e.g. Chelex), 2) organic extraction (e.g. Phenol/PCI), 3) differential precipitation (e.g. salting out the DNA), or 4) differential affinity (e.g. silica). \par \par \par }{\b\f7 \tab A. Extracting from skins:}{\f7 \par \par Stock Chemicals & Solutions: \par \par Amresco makes pre-made stocks at semi-reasonable prices. I used them & had good luck. I would recommend pre-made stocks to anyone not working with ancient human stuff. \par \par }\pard \li360\sl240\slmult0\widctlpar\adjustright {\f7 EDTA - Amresco # E177 \par Tris - Amresco # E199 \par SDS - Amresco # 0837 \par PCI - Phenol Chloroform Isoamyl Alcohol (25:24:1), Amresco # }{0883}{\f7 \par CI - Chloroform Isoamyl Alcohol (24:1), Amresco # }{0883}{\f7 \par Proteinase K - Amresco # }{E195}{\f7 \par DTT - powder, Amresco # 0281 \par CaCl}{\f7\fs20\sub 2}{\f7 - Amresco # }{0556}{\f7 \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par }\pard \li360\sl240\slmult0\widctlpar\adjustright {\f7 Collagenase - Sigma Type I, # C-0130 \par NaCl - \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par Materials: \par \par }\pard \li360\sl240\slmult0\widctlpar\adjustright {\f7 Microcon 30's - Amicon # 42410 \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par Solutions: \par \par }\pard \sl240\slmult0\widctlpar\tx3780\adjustright {\f7\ul Components}{\f7 \tab }{\f7\ul Stock to add per mL}{\f7 \par }\pard \sl240\slmult0\widctlpar\tqdec\tx980\tqdec\tx4320\adjustright {\f7 \par TE:\tab 10 mM Tris pH 8\tab 10 \u181\'b5L of 1M \par \tab 2 mM EDTA\tab 4 \u181\'b5L of 0.5M \par \par TLE:\tab 10 mM Tris pH 8\tab 10 \u181\'b5L of 1M \par \tab 0.2 mM EDTA\tab 0.4 \u181\'b5L of 0.5M \par \par P\u228\'8a\u228\'8abo et al. (1988) Digest Buffer: \par \par }\pard \sl240\slmult0\widctlpar\tqdec\tx540\tqdec\tx4320\adjustright {\f7 \tab 10 mM Tris pH8\tab 10 \u181\'b5L of 1 M \par \tab 2 mM EDTA\tab 4 \u181\'b5L of 0.5 M \par \tab 10 mg/mL DTT\tab 100 \u181\'b5L of 100 mg/mL \par \tab 0.5 mg/mL Proteinase K\tab 50 \u181\'b5L of 20 mg/mL \par \tab 0.1% SDS\tab 10 \u181\'b5L of 20% \par x dH}{\f7\fs20\sub 2}{\f7 O\tab 826 \u181\'b5L \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par TCG Digest Buffer: \par \par }\pard \sl240\slmult0\widctlpar\tqdec\tx540\tqdec\tx4320\adjustright {\f7 \tab 12.5 mM Tris pH8\tab 12.5 \u181\'b5L of 1 M \par \tab 12.5 mM NaCl\tab 12.5 \u181\'b5L of 1 M \par \tab 1.25 mM CaCl}{\f7\sub 2}{\f7 \tab 12.5 \u181\'b5L of 100 mM \par x dH}{\f7\fs20\sub 2}{\f7 O\tab 962.5 \u181\'b5L \par \par to 800 \u181\'b5L of the above, you will add: \par \tab 0.5 mg/mL Collagenase\tab 50 \u181\'b5L of 20 mg/mL \par then: \par \tab 0.5 mg/mL Proteinase K\tab 50 \u181\'b5L of 20 mg/mL \par \tab 10 mg/mL DTT\tab 100 \u181\'b5L of 100 mg/mL \par \tab 1% SDS\tab 10 \u181\'b5L of 20% \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 and finally: \par }\pard \sl240\slmult0\widctlpar\tqdec\tx540\tqdec\tx4320\adjustright {\f7 \tab 0.5 mg/mL Proteinase K\tab 50 \u181\'b5L of 20 mg/mL \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par PCI should be equilibrated to pH 8 (for Amresco PCI, simply dump in the pre-measured Tris). \par \par }{\f7\ul Procedure}{\f7 \par \par Note: you may use the P\u228\'8a\u228\'8a bo et al. (1988) Digest Buffer rather than the TCG Digestion Buffer. Theoretically the proteinase K is about 10x more active in the TCG digestion buffer. In practice the TCG buffer provides somewhat better digestion, but it isn't that dramatic. \par \par }\pard \fi-440\li440\sl240\slmult0\widctlpar\adjustright {\f7 1) Put on gloves and clean a surface to work on, wiping down with 10% bleach. If possible, work on a clean Kimwipe. \par \par 2) Place 2+ forceps into beaker with bleach diluted 1:1 with dH}{\f7\fs20\sub 2}{\f7 0. Fill another beaker with dH}{\f7\fs20\sub 2}{\f7 0. Note: the forceps must be in bleach for }{\b\f7 at least 5 min.}{\f7 to destroy DNA to \u8804\'b2 50 bp. If you work quickly, you will want to have enough forceps to ensure that each pair soaks for at least 5 min. \par \par 3) Get out a specimen (AKA old dead bird tissue). \par \par 4) Record specimen number & data on record sheet. \par \par 5) Write specimen number on a 1.7 mL tube and add 400 \u181\'b5L of Digestion Buffer to the tube. \par \par 6) Place one set of forceps into beaker of dH}{\f7\fs20\sub 2}{\f7 O. \par \par 7) Clean forceps with kimwipe & set aside. \par \par 8) Open new disposable scapel & break-off blade guard. \par \par 9) Grasp the desired piece of skin with the forceps and place it in a new plastic weigh boat. \par \par 10) Cut a piece - about 0.25 cm x 1 cm - (a sub-sample) from the original sample. Cut off attached feathers at the base (leaving the quill in the skin). \par \par 11) Return the rest of the sample to the ziplock baggie. \par \par 12) Squirt 400 \u181\'b5L of Digestion Buffer onto the subsample. \par \par 13) Cut the subsample into as many small pieces (shavings) as possible. Be careful not to cut through the weigh boat. \par \par 14) Transfer the subsample and buffer to the 1.7 mL tube. \par \par 15) Place the second set of forceps into the dH}{\f7\fs20\sub 2}{\f7 0. \par \par 16) Wipe-off the first set of forceps (the ones you just used) with a kimwipe and then place them into a beaker with diluted bleach. \par \par 17) Toss the weigh boat & Kimwipes. \par \par 18) Rewrap the scapel inside its original foil (for disposal or later use with modern samples). \par \par 19) Return the specimen to the appropriate drawer. \par \par 20) Change gloves if you touched any samples. Replace the Kimwipe that you are working on. \par \par 21) Repeat steps 3 - 20 for all remaining specimens, unless you plan to process more than about 12. If working with \u8805\'b312 samples, group samples into appropriate sets & proceed to the next section when you have completed all samples in a set. \par \par 22) Complete a negative control for the set by doing all manipulations as above, but without any tissue (i.e. place 400 \u181\'b5L of digestion buffer into a tube, squirt another 400 \u181\'b5 L into a weigh boat, slosh a pair of forceps in the weigh boat & transfer the buffer to the tube). \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par }\pard \sl240\slmult0\widctlpar\tqdec\tx540\tqdec\tx4320\adjustright {\f7 23) to 800 \u181\'b5L of the above, add 50 \u181\'b5L of 20 mg/mL Collagenase (0.5 mg/mL final concentration). \par \par 24) Incubate at 37}{\f4 \u730\'fb}{\f7 C for 1 to 3 hours. If you have additional sets, then start on the next one \par \par 25) Add: \par \tab 0.5 mg/mL Proteinase K\tab 50 \u181\'b5L of 20 mg/mL \par \tab 10 mg/mL DTT\tab 100 \u181\'b5L of 100 mg/mL \par \tab 1% SDS\tab 10 \u181\'b5L of 20% \par }\pard \sl240\slmult0\widctlpar\adjustright {\f7 \par 26) Incubate at 55}{\f4 \u730\'fb}{\f7 C overnight, while shaking or rocking. \par \par 27) To samples that still have intact tissues, add another 50 \u181\'b5L of 20 mg/mL Proteinase K (1 mg/mL final concentration). \par \par 28) Incubate at 55}{\f4 \u730\'fb}{\f7 C, while shaking or rocking, for an additional 1 to 3 days. \par \par 29) Extract following the Phenol Chloroform Extraction procedure below. \par \par \par }{\b\f7 Phenol Chloroform Extraction \par }{\f7 \par Note: PCI & CI should always be handled in a functional chemical fume hood. Thus, all steps that involve open tubes that contain PCI or CI, should be done in a hood. \par \par 1) Remove tubes from 55o oven & spin 5 seconds at full speed to precipate large particles. \par \par 2) Label a new 1.5 mL tube for each sample. \par \par 3) Add 5 \u181\'b5L of 0.2 M }{\b\f7 EGTA}{\f7 to the new tubes (or 2 \u181\'b5L of 0.5 M EDTA). \par \par 4) Transfer 500 \u181\'b5L of supernatant from each sample to the appropriate new tube. \par \par At this point, it is wise to group the samples into sets, with 4-8 samples in each set. I prefer to process two sets of 6 samples as described below. \par \par 5) Add 500 \u181\'b5L of PCI to each tube in the first set. Shake vigorously, or vortex each tube. \par \par 6) Place the first set of samples in the IEC microcentrifuge & spin at full speed for ~5 min.. \par \par While this set is spinning, another PCI can be added to another set. \par \par 7) Remove the first set from the centrifuge & place the next set in. Spin the second set. \par \par While the second set is spinning: \par \par 8) Transfer the aqueous (top) layer above the PCI to a new tube. Avoid taking particles from the PCI interface. \par \par 9) Add 500 \u181\'b5L of CI to each tube with aqueous sample. Shake vigorously, or vortex each tube. \par \par 10) Remove the second set of PCI samples & place the CI samples into the microfuge. \par \par 11) Spin the tubes at full speed for 3+ min. in the IEC microcentrifuge. \par \par 12) Repeat steps 8 & 9 for the second set of tubes. \par \par 13) Remove the first set of CI samples from the microfuge & place the second set in it. \par \par 14) Spin the second set of CI samples at full speed for 3+ min. in the IEC microcentrifuge. \par \par 15) Transfer the aqueous (top) layer above the CI to a labeled microcon tube. Avoid taking particles from the CI interface. \par \par 16) Repeat step 15 for the second set of samples. \par \par 17) Spin the Microcon tubes for 10 min. at 12,000 RPM in the STE microcentrifuge. \par \par 18) Remove the Microcon tubes from the STE. Carefully pull out the concentration cup, dump the filtrate into a waste beaker*, and replace the conce ntration cup. [* the filtrate can be saved in a new tube, if you wish to characterize it, &/or to ensure that it doesn't have the DNA] \par \par 19) Add 400 \u181\'b5L of TLE to the Microcon tubes. \par \par 20) Spin the Microcon tubes for 8 min. in the STE microcentrifuge. \par \par 21) Recover the retentate TLE (& DNA) by placing the concentration cups upside down in a new labeled Microcon collection tube. Spin in the IEC microcentrifuge for 15 seconds. \par \par 22) Return the concentration cups to the original collection bottoms. \par \par 23) For samples where <75 \u181\'b5L of retentate is recovered, add enough TLE to the concentration cup to bring the volume to ~75\u181\'b5L (e.g., if 40 \u181\'b5L is recovered, then add 30 \u181\'b5L to the concentration cup) & repeat steps 21 & 22. \par \par 24) Characterize recovered DNA by PCR. I usually keep the filters and filtrate. I toss them later, when I realize that I will never have the time to characterize the amount of DNA in/on them. \par \par }{\b\fs28 \page Chelex DNA Preparation Protocol for "Ancient Tissue" \par \par Overview \par }\pard \li260\ri-80\sl240\slmult0\widctlpar\adjustright { Based on the protocol of: Walsh, S. P., D. A. Metzger, and R. Higuchi. 1991. Chelex 100 as a medium for simple extraction of DNA for PCR-based typing from forensic material. Biotechniques 10(4): 506-513. See also Morin, P. A. and D. S. Woodruff. 1992. Paternity exclusion using multiple hypervariable microsatellite loci amplified from nuclear DNA of hair cells. Pages 63-81 in R. D. Martin, A. F. Dixon, and E. J. Wickings (eds). }{\ul Paternity in primates: genetic tests and theories.}{ Basel, Karger. Chelex 100 is available from Biorad (1-800-4BIORAD, catalog # 143-2832 -> $75/100g = <$0.01 per "extraction"). \par }\pard \sl240\slmult0\widctlpar\adjustright {\b\fs28 \par }{\b\f26 Preparation of Chelex 100 Master Mix and Aliquots \par }{\f5 \par Adapted from Museum of Vertebrate Zoology (MVZ) protocol. \par \par \par 1) Obtain a new 50 mL polyethylene (Falcon) conical tube. \par \par 2) Sterilize a spatula and the tiniest magnetic stir-bead by soaking in a beaker of Chlorox for 10}{\f5\super +}{\f5 min. After soaking, rinse the stir-bead and spatula thoroughly with dH}{\f5\fs20\sub 2}{\f5 O. Note: the stir-bead can be held in place by using a larger stir-magnet on the outside of the beaker. \par \par 3) Place the stir-bead in the conical tube. Then, place the conical tube + stir-bead on the scale, and weigh 2.0 grams of Chelex 100 into the tube. \par \par 4) Fill the conical tube to the 40 mL mark with dH}{\f5\fs20\sub 2}{\f5 0. \par \par 5) Place the conical tube on a stir plate and let the stir-bead do its thing. Ensure that the entire volume of Chelex + dH}{\f5\fs20\sub 2}{\f5 0 is stirring. Note: You may wish to do this step and the next one in a Laminar Flow Hood. \par \par 6) Using the P1000 and filtered pipette tips, aliquot the stirring 5% Chelex into 1.7 mL micro-centrifuge tubes obtained from a freshly opened bag. It is usually best to make both 500 \u181\'b5L aliquots and 200 \u181\'b5L aliquots.}{ \par \par \par }{\ul Procedure}{ \par \par 1) Place Eppi-pestels, forceps, scalpel handles, a small container for liquid Nitrogen, etc. in a solution of Chlorox for 20}{\super +}{ minutes. (Hydrochloric acid may also be used here.) \par \par 2) Rinse Eppi-pestels, etc. with clean dH}{\fs18\sub 2}{O and expose them to UV for 10+ minutes. \par \par 3) Obtain CLEAN 5% Chelex tubes from the rack in the kelvinator (prepared using MVZ protocol), and turn on the heating block. \par \par 4) Aliquot 10-20 ul of the aqueous Chelex solution from the original tube(s) into a clean 600 ul tube (each Chelex tube will have its own separate aliquot). \par \par 6) Prepare tiny portions of the tissue to be ground-up and Chelexed. One milligram of tissue is about the right amount. Place the tissue pieces in separate tubes. \par \par 5) Place the liquid Nitrogen into the clean container for it. \par \par 6) Dip a Clean 1.5 ml eppi-tube into the liquid Nitrogen. \par \par 7) Quickly place a TINY amount of tissue into the tube when it is about 1/2 full of liquid Nitrogen (using your forceps). \par \par 8) Grind the tissue using the Eppi-pestel. \par \par 9) Transfer the Chelex solution to the tube where you have ground-up the tissue. Pipette up and down several times to mix the chelex solution and the tissues. \par \par 10) Incubate in the heating block at 95-100 C for 10 minutes to however long it takes to do step 11. \par \par 11) Prepare your PCR master mix. You will be need enough for each Chelex tube, the aliquot from each Chelex tube, a positive control, and a negative mix control. \par \par 12) Take your Chelex tubes off the heating block. \par \par 13) Distribute the PCR mix into the PCR sample tubes. \par \par 14) Place 2 ul of Chelex solution (}{\b Avoid the Beads}{) into the sample tubes. You should physically stir by swirling the pipette tip in the upper aqueous portion of the tube, and pumping up and down. This will ensure that DNA in the tube is in the aqueous solution, but doesn't seem to stir up the beads too much. \par \par 15) Thermocycle with normal parameters. (You will often find it necessary to add 5 cylcles and lower the annealing temperature 5 degrees to get satisfactory amounts of product DNA because you start with very few copies, and the DNA is often damaged or modified in a funky way.) \par \par \par Note: If you do not get amplification, try taking 20-100 ul of your original Chelex solution, and place it into a new Chelex (i.e. dilute it). This ha s often resulted in amplication when other methods have failed. The major problem encountered is using a small enough piece of tissue, and a small enough target fragment. If you continue to have problems, go for a smaller fragment (I recommend 100-150 b p ) and test the dilution of your ancient DNA solution that will kill PCR of good modern DNA. Other magic bullets include: BSA, 2-100 ug/ml ; DMSO, 10%; Glycerol, 10%; single-strand binding protein; and Gel or Column purifications (many methods will w ork - we are still trying some of these). \par }{\f7 \par }{ \par }}