Step by Step SSCP

Travis Glenn
Laboratory of Molecular Systematics
Smithsonian Institution
Washington, DC  20560
phone:  301-238-3444
fax:  301-238-3059
e-mail:  Glenn@onyx.si.edu

Warning:  This protocol is currently being updated.  It is dangerous to use 35S in PCR reactions.  It forms a volatile gas that WILL escape from the PCR tubes (see Clinton & Scougall 1995).

	Below I present the protocol for Single-Strand Conformation Polymorphism (SSCP) analysis used at LMS.  This protocol is basically the same as any SSCP protocol you will find in the literature.  The only major difference is that I recommend doing a two reaction PCR (unless you are screening more than 48 individuals, then you should optimize PCR conditions and amplify hot the first time around).  The first reaction is a “normal” cold PCR to amplify the intended products.  Check that these reactions were successful by running out on a minigel.  The second reaction is a hot PCR to label the products produced in the first PCR reaction.  The hot product will be diluted in sequencing buffer, denatured, and loaded onto a native polyacrylamide gel.

	I have structured the protocol with 4 major tasks.  Within each task, I have included a step by step procedure to accomplish the task.  At the beginning of each task I have included one or two times (e.g., task 3 (~3 hrs. => ~7 hrs.)).  The first time listed (~3 hrs) is the approximate amount of "hands on" time that task will require.  A second time is listed following a => if the procedure includes "hands off" times (e.g., incubations).  The second time is the total time required to complete the task.

	The primary purpose of this protocol is to serve as a guide for people at the LMS (in the spirit of Klone Alone).  A secondary purpose of this protocol is to spread the knowledge I (we) have accumulated at the LMS (your tax dollars at work).  Because of the diverse nature of the people working at the LMS, I intend this protocol to be most useful to those who are inexperienced.  Therefore, I have included lots of details that most experienced workers will find quite boring & unnecessary.  If you are familiar with a procedure, then read enough to get the gist, and do the task in the way you are familiar with.

There are, no doubt, still many errors which remain to be caught.  If you find one, please let me know so I can fix it.

Thanks and Good Luck.

Travis Glenn  22 March 1996.

SSCP Background Information

	Single-Strand Conformation Polymorphism analysis was orginally described by Orita et al. (1989).  The general idea is to take a small PCR product (a.k.a. amplicon), denature it, and electrophorese it through a non-denaturing polyacrylamide gel.  Thus, as the PCR product moves into and through the gel (and away from the denaturant), it will regain secondary structure that is sequence dependent (similar to RNA secondary structure).  The mobility of the single-stranded PCR products will depend upon their secondary structure.  Therefore, PCR products that contain substitutional sequence differences as well as insertions and deletions will have different mobilities.

	The major advantage of SSCP is that many individual PCR products may be screened for variation simultaneously.  Forty nine to 97 PCR products may be analyzed on each full-size sequencing gel (depending upon comb size used).  Most researchers use SSCP to reduce the amount of sequencing necessary to detect new alleles at loci of interest (e.g. Sweetman et al. 1992) or to better estimate allele frequencies of populations (e.g. Aguadé et al. 1994).  

	I advocate two additional uses of SSCP.  First, SSCP should be used to screen PCR products of genes that researchers intend to sequence for phylogenetic analysis.  This SSCP screening will allow researchers to determine 1) if the gene contains sufficient polymorphism that sequencing will be worthwhile, 2) which portion of the gene is most polymorphic, 3) what level of intraspecific variation exists, and 4) if there is polymorphism among multicopy genes within individuals (e.g. rDNA).  Second, SSCP is a much more efficient method of obtaining information about levels of polymorphisms within anonymous nuclear loci than the restriction enzyme protocol originally described (Karl et al. 1992).  The advantages of using SSCP in this context are:   1) the PCR amplicons are smaller and thus easier to amplify,  2) because the amplicons are only 200-300 bp, the entire sequence needed for PCR primer design can be obtained from a single sequencing reaction,  3) similar amounts of information are obtained by simply running the amplicons on 2 SSCP gels (e.g. one with glycerol and one without), rather than doing 50 restriction digests and assumming that the restriction sites are independent (a problematic assumption when dealing with a 1 kb amplicon), and 4) when polymorphisms are detected, the amplicons are of optimal size to sequence using an ABI automated sequencer (so getting sequence level information is easy).

	There are 2 major disadvantages of SSCP.  First, the amount of mobility differences have little if any correlation to the amount of sequence differences.  For example, under some SSCP conditions, cytochrome b PCR products from one subspecies of Red Panda are closer in mobility to Raccoons than to the other subspecies of Red Panda.  Thus, the only information that can be gained from SSCP is if PCR amplicons are “identical” or different.  Second, the optimal amplicon size for detection of most point mutations is rather small, around 200 bp.  The strategies to deal with this limitation (e.g. dideoxy fingerprinting or cutting amplicons with restriction enzymes) are often more trouble than they are worth.

	Other methods of quickly screening amplicons have been described (e.g. Kirkpatrick et al. 1993).  For a general review, I highly recommend Lessa and Applebaum (1993; see also Langemeier et al 1994 and references therein for dideoxy fingerprinting).  Lessa and Applebaum recommend Denaturing Gradient Gel Electrophoresis (DGGE) as the most widely useful method.  I agree, that with their research goals, DGGE is going to perform better than SSCP most of the time.  The essential trade-offs between DGGE and SSCP are:  1)  DGGE can detect single-base substitions more efficiently in longer amplicons (about twice the optimal length for SSCP),  2)  DGGE requires that one of the PCR primers include a "GC-clamp" (about 40 bp of G's and C's; which is about as expensive as getting 2 additional primers to either screen a second fragment of the same locus or get another locus -> thus primer costs per bp screened are similar),  3)  DGGE requires one to determine the optimal gradient conditions before screening individuals for variation,  4)  DGGE requires additional specialized equipment.  Therefore, I feel that when one is concentrating on finding intraspecific variation within one genus (insects) or family (vertebrates) of organisms, a small number of primers and conditions will be highly useful and thus DGGE is the method of choice.  However, when one wants to screen many loci or the organisms of interest are quite dissimilar at the sequence level (e.g. raccoons vs. red pandas) then SSCP will usually be better.

	Finally, many methods of screening for SSCP variants have been described.  For a general review, I recommend Sheffield et al. (1993) and Vidal-Puig and Moller (1994).  Both ethidium bromide (Yap and McGee 1993) and silver-staining can be used, rather than radiation to detect SSCP alleles.  I would not recommend ethidium bromide.  Silver staining works, but is not as reliable (for me) as using radiation.  If I were going to do silver staining, I would probably use a mid-size gel rig (e.g. Protean II from Stratagene) and the DP-SSCP suggested by Yap and McGee (1993).  Also, one can use "normal" polyacrylamide rather than MDE or Long Ranger (from AT Biochem).  I prefer Long Ranger and MDE because they never stick to the glass rather than blotting paper, thus I never have to fix my gels.  So far, I have not used glycerol in my SSCP gels.  I plan to do this in the next couple of months, and will add this to the protocol after I get it to work well.

Acknowledgments

	Most of the gel pouring information is adapted or directly taken from Caleb Gordon's Sequence Gel Protocol (for BRL [S2] gel rigs).

I.  Initial PCR Reaction

1)  Set-up a “normal” cold PCR reaction.  The following recipe is generally used:

			Master Mix					
	2.5 µL 10x Thermo Buffer
	2.5 µL Primer 1 (5 µM)
	2.5 µL  Primer 2 (5 µM)
	1.5 µL MgCl2 (25 mM)
	1.5 µL dNTPs (2.5 mM each)
	11.8 µL dH2O
	0.2 µL Taq DNA Polymerase (1 unit)	

	Multiply each component by the number of reactions you plan to do + 1 for a negative control + 10% (e.g. to do nine experimental reactions - multiply each component by 11, so the master mix would contain 27.5 µL 10x Thermo Buffer, ...).  ENSURE THAT EACH COMPONENT IS THOROUGHLY MIXED BEFORE ADDING IT TO THE MASTER MIX.  Aliquot 22.5 µL of the Master Mix into a tube for each reaction.  Then, add 2.5 µL DNA (@20 ng/µL) to each tube, ENSURING THAT THE DNA IS THOROUGHLY MIXED.  Add 1 drop of mineral oil to each reaction tube (unless you plan to use the hot lid - Cycler-Mate, BioLogic Engineering, Inc.).  If you plan to use the hot lid when screening your samples, then you must use it while optimizing the PCR conditions.  
2)  Thermocycle with the parameters established during optimization.  In general, the following parameters are used:  1 cycle of 94o for 2 min., then 25 to 30 cycles of 94o for 1 min, 50o* for 30 sec., 72o for 30** sec.  [Note:  *50o is a standard annealing temperature, use temperature established during optimization; ** 30 seconds is usually sufficient to synthesize PCR products less than 400 bp, use 1 min. per kb if synthesizing longer products.]  
3)  Examine your results on a 1.5% agarose minigel.  Use Gel Marker I (Research Genetics, Huntsville, AL) mixed with Light Loading Buffer (NOT the buffer that comes with the Gel Marker I) as a size marker.  Use light loading buffer to screen 10 µL of each sample.
4)  If the reactions contain anything other than bands of approximately the correct size, try the following to re-optimize conditions:
	1)  Change the annealing temperature
	2)  Add 2.5 µL of BSA (2.5 mg/mL - Non-acetylated)
	3)  Increase the MgCl2 concentration 2x or 3x
	4)  Add 2.5 µL of DMSO
	5)  Add 2.5 µL of Glycerol

[Note:  if you add anything to the PCR reaction (e.g. BSA) be sure to subtract an equal amount of water, so the total reaction volume remains constant.]

II.  Making the SSCP Gel

Note:  Always be very careful when handling acrylamide.  It is a potent neurotoxin.  Use gloves, glasses and lab coat when handling

A) Gel Mix (15 - 30 min.)
1)  Make gel solution:
For one MDE gel:	For five gels:
15 mL   2 x MDE	75 mL 2 x MDE
3.6 mL 10 x TBE	18 mL 10 x TBE
41.1 mL dH20	205.5 mL dH20

For Gels with 10% Glycerol:
For one gel:	For five gels:
16.5 mL   2 x MDE	82.5 mL 2 x MDE
4 mL 10 x TBE	20 mL 10 x TBE
6.6 mL Glycerol	33 mL Glycerol
38.7 mL dH20	193.5 mL dH20

For Glycerol Gels, it is usually convenient to measure the glycerol in a 25 mL graduated cylindar, then fill the cylindar with dH20 (18.4 mL dH20), cover with parafilm, and mix by inversion.  This way, the glycerol is diluted & easier to manipulate.  The cylindar is then "washed with 20.3 mL dH20.
If making enough for multiple gels, then store the mix in a brown plasic bottle.  The mix should be used within 1 month of being made.
Note:  Each MDE gel will also contain 330 µL APS + 30 µL TEMED and 10% Glycerol gels will contain 400 µL APS + 36 µL TEMED added immediately prior to pouring the gel (see below).
2)  When making enough mix for multiple gel, it is usually a good idea to filter the Gel Mix.  This should be done with the reusable magnetic filter units and the 0.22 micron AcetatePlus filters (MSI cat. # A02SP04700).  
3)  Get frozen aliquot of 10% Ammonium Persulfate (APS) or make fresh 10% APS solution by adding 1 gram of APS to a 14mL snap cap tube and then filling to 10 mL with dH20.  [Note:  the excess 10% APS can be aliquoted and frozen for future use.]

B)  Setting up and Pouring a SSCP Gel (30min.-1hour)
Setting up the gel mold (glass plate sandwich):
1)  Take one long glass plate and one short glass plate from the drying racks to the Acrylamide Gel Pouring Area.
2)  Place the plates on the table with the inside surfaces of the plates facing up (inside surface can be identified by etched notation on outside surface of plate).  Treat the short plate only with rainex, then spray and wipe dry both plates with deionized water, and then 95% ethanol.
3)  Make "glass plate sandwich” which rests on two empty pipette racks by first placing long plate on pipette racks inside surface facing up.  Then wipe two plastic spacer strips with ethanol and place one along each of the long edges of the long plate with the rubber cushions at the top and facing up.  Place the short plate directly on top with the inside surface down and make sure the plates and spacers are lined up precisely with the rubber cushions of the spacer strips fitting snugly against the top edge of the short plate.
4)  Place three clamps along each long side of glass plate sandwich leaving about 6 inches without clamps at the bottom.  Pressure points of clamps should be directly over the center of the spacers.  
5)  OPTIONAL:  insert a strip of 3MM filter paper 1 cm thick into the bottom of the gel sandwich.
6)  Carefully place one long strip of sequence gel tape along the bottom edge of the sandwich and coming about 5 inches up the side edges.  Make sure the tape is sealed tightly along the bottom and side by firmly pressing it along the edges of the glass (the depth) before folding it and sealing along the length & then width of the glass.
7)  Put a fourth clamp on each side of the sandwich near the bottom (on the taped area).

Adding polymerizing agents and pouring gel (to be done only when gel mold is ready)
8)  Ensure the following items are ready to use:  60cc syringe; 18 gauge, 1.5 inch needle; 100 mL graduated cylinder; 150 mL beaker; P1000 pipetter set to 330 µL; P200 pipetter set to 30 µL; 10% APS; TEMED; gel mix; and gel mold
9)  Measure out 55-60 mL of gel mix.  Add 330 µL of 10% APS and then 30µL of TEMED to the gel mix .
	note:  a minimum of 45 ml is enough for a gel but it is a good idea to have at least some extra in case of some drippage and to monitor polymerization.
10)  Using 60cc syringe without needle, suck up solution.  Wipe the end of the syringe and put the 18 Guage 1.5 inch needle on.
11)  Pick up gel mold and tilt it up (45 degree angle with table or greater) and to one side (slightly).  Begin squirting gel mix into the opening between the plates at a point near the low side edge (near the spacer cushion). 
12)  Squirt in gel mix slowly and steadily.  Try to avoid forming bubbles or spilling.  Spilling can be avoided by squirting slowly.  Bubbles can usually be avoided by increasing the angle of tilt to the side or more vertically as you pour.
13)  When the fluid is about three inches from reaching the top of the mold, stop squirting and place the mold, long plate down, flat on the empty pipette racks again.  The fluid should come up to fill the entire gap between the plates.  Squirt remainder of fluid back into bottom of the 150 mL beaker so that polymerization may be monitored.
14)  Slide the two plastic combs in between the plates into the gel mix with the flat sides in (teeth out).  Flat surfaces of combs should form an even line, about a half centimeter deep into the gel (the top of the oblong holes should be even with the top of the short glass plate).
15)  Place a large piece of saran wrap around top of gel mold and avoid disturbing combs.
16)  Take one clamp from each side of the gel mold and place them clamps along top of gel mold.  Spread the 3 remaining clamps along each side evenly.
17)  Let gel sit on empty pipette racks for a minimum of an hour (up to overnight)

III.  Preparing and Loading Samples

I generally start here after lunch (around 2:00 pm).  If you skipped step one, then increase the number of cycles to the number determined during primer optimization and ensure that the MgCl2 concentration is the same as determined during optimization rather than what is given below (it may also be ok to double the dNTP concentration).

A)  Making "Hot" PCR amplicons (1 hour => 2 hours)

1)  Label one tube for each PCR product.  Then make the following:

The New Less Dangerous Way

1. A.  End label one of the primers using the following recipe (enough for 50 individuals/reactions):

	5.0µL 10x Kinase Buffer
	20.0 µL Primer (5 µM)
	20.0 µL dH2O
	2.5 µL PNK (PolyNucleotide Kinase - Promega - 8 units/µL)
	2.5 µL gamma 33PdATP (10 mCi/mL; 2000 Ci/mmol = 5 pmol/µL)

Incubate at 37o for 30+ min.

1. B.  Label the PCR products with the hot primer by making new reactions according to the following recipe:

		Master Mix					
	1.00 µL 10x Thermo Buffer
	0.75 µL Hot Primer (2 µM)
	0.30 µL Cold Primer (5 µM)
	0.6 µL MgCl2 (25 mM)
	0.6 µL dNTPs (2.5 mM each)
	5.65 µL dH2O
	0.1 µL Taq DNA Polymerase (0.5 unit)	

The Old Dangerous Old Way:

			Master Mix					
	1.0 µL 10x Thermo Buffer
	1.0 µL Primer 1 (5 µM)
	1.0 µL Primer 2 (5 µM)
	0.6  µL MgCl2 (25 mM)
	0.3 µL dNTPs (2.5 mM each)
	3.5 µL dH2O
	0.5 µL  35S dATP  (Note: use alpha labelled nucleotide)
	0.1 µL Taq DNA Polymerase (0.5 unit)	

Handle all radioactive materials with much care to avoid contamination.  Monitor everything frequently to avoid spreading radiation.

2)  Aliquot 8 µL of master mix to each reaction tube.  (Mix well by gently pumping pipette before making aliquots!)  
4)  Add 2 µL of previous PCR products (or DNA if you did not do step I) to each reaction tube.
5)  Add 1 drop of mineral oil to each tube and then close.
6)  Using the same thermocycling parameters as used in step I, cycle 10-15 times (the intial denaturation isn’t necessary). 
7)  Turn on the radioactive-use water bath to 90o. 

B)  Preparing Samples for Loading into SSCP Gel (30 min)
1)  While the thermocycler is still running, add 18 µL of SSCP Loading Buffer (USB Stop Solution + 2 µL of 10 M NaOH per mL) to 0.7 mL tubes (one for each reaction done in part III A).  [Note:  the P250 electronic pipetters are great for this.]  If you have many samples, it is easier to use a microtiter plate here, rather than tubes.  [Note:  adding 1 µL of 1 M CH3HgOH per sample will ensure denaturation - see Weghorst & Buzard (1993)].
2)  Label the tubes with SSCP Loading Buffer, with the appropriate sample numbers.  Centrifuge the tubes to ensure the drops of Loading Buffer are on the bottom.
2)  After the amplicons have been labelled (the thermocycler is done cycling), add 2 µL of the Hot amplicon to the appropriate tube made in step 1 (above).  

C)  Preparing and Loading SSCP Gel (1hour)
I generally am here at about 4:30 or 5:00.  It is best not to start too early, because the gel will run 14-18 hours (i.e. overnight).
1)  Make sure radioactive-use water bath is at 90-95o C.
2)  Check small sample of gel mix (remainder from step B15 above) to make sure that gel polymerized correctly.  If it is still mostly liquid, there may have been a problem with the acrylamide mix or one of the polymerizing reagents and you should not waste your reactions on the gel; make a new gel mix, NEW APS & pour again.
3)  Remove the tape from the bottom, then remove the saran wrap, then all the clamps from gel mold, and finally, take out combs carefully to avoid damaging top edge of gel.
4)  Place gel, long-plate-out, on gel rig and tighten clamps to secure gel.
5)  Close buffer drain valve by turning knob on right side of rig clockwise until tight.
6)  Get 1 liter of 1x TBE** from carboy, and pour enough into bottom tray of gel rig to cover bottom of gel (cover about 1 cm of the bottom of the glass plates).  Pour the remainder in the top compartment of the gel rig.
**  The concentration of TBE may be varied in this step.
7)  Suck up some of the buffer, and squirt all along top edge of gel to remove air bubbles and gunk.  You may use a Pasteur pipette, or a syringe (with needle) for this step.
8)  Shut plastic lids on top and bottom compartments of gel rig and use electrical cords to connect gel rig with power source (make sure positive charge is at the bottom of the rig).
9)  Turn on power source and set to desired Wattage (usually 4-6).  Run gel for about 5 minutes before loading to ensure that it will carry current and there are no huge leaks.
10)  Float  reactions (either microtiter plate or eppendorf tubes) in 90o water bath for 2 minutes to denature the amplicons.  If using microtiter plates, make sure water contacts the bottoms of all of the wells with reactions in them.
11)  While reactions are incubating, get ice bucket ready and bring the following materials to the gel rig:  magic marker, P10 pipette man, P10 pipette tips, 60cc syringe with needle or Pasteur pipette with bulb, and a labelled beaker for temporary storage of used radioactive pipette tips.
12)  After removing reactions from water bath, immediately place them on the ice.
13)  Turn off electrical current on rig, open lid on top compartment and squirt buffer along the top of the gel to ensure that all the gunk is rinsed out.
14)  Carefully slide toothed side of plastic combs into top of gel such that the tips of the teeth protrude only slightly into the gel (roughly 0.5-1mm).  If comb slides in too easily, the wells will leak, so keep trying combs until you get two that fit snugly.
15)  Using magic marker, mark lanes to be used.  Select lanes with no bubbles and with a good contact with teeth.  Samples should be run side-by-side as much as possible.  It will be very difficult to judge differences if there are gaps between the samples.
16)  Load 1-3 µL of each reaction into wells if using fine toothed combs, 3-5 µL if using larger toothed combs.  
17)  When all lanes are loaded, close lid, reconnect power source, and turn on power.
18)  Mark your name, time of the start of the run, and times to stop the gel on the outside of the gel with a magic marker.  Also mark a line at the level of the buffer near the top of the glass plate.  This step will allow you to monitor the rig for buffer leaks.  If you notice buffer leaking from the top compartment of the gel rig at this point or at any time during the run, disconnect the power and take steps to plug leak.  Using two syringes with needles to poke a small piece of kimwipe into the corner where the rubber gasket touches the glass plate usually does the trick.
19)  Make sure that the order of the samples, the primers, templates, reaction conditions and order of lanes on the gel are all written down on a data sheet.  The left-right orientation of the gel can easily get confused in later gel handling steps.  A good way to eliminate this confusion is to load the lanes in an asymmetrical configuration (e.g. 1,2,3,4,5,   6,7,8).  
20)  There is a fair amount of variation in the speed that individual gels run.  For MDE gels, I have had good luck running gels at 4 to 8 Watts for about 40 Watt hours (= 10 hours at 4 Watts or 5 hours at 8 Watts, or ... ) when amplicons are 50 - 100 bp;  50-60 Watt hours with 100 - 200 bp amplicons; etc.  Glycerol gels take considerably longer to run (usually a bit less than 50% longer).  The cyanol blue (light green dye) usually runs at about 100 bp.  Use that as a guide.

IV)  Taking off, drying and exposing gel

A)  Taking gel off rig (15 minutes)
1)  Turn off power source and disconnect wires.
2)  Unscrew buffer valve on right side of rig to allow buffer in top compartment to drain into bottom compartment.
3)  When buffer is drained out of top, unscrew clamps, remove gel from rig and place, long-plate-down, on a flat surface with bench paper.
4)  Allow gel to cool slightly while you remove buffer tray from bottom of gel rig and dump buffer from both compartments into radioactive liquid disposal carboy in sink (contains radiolabelled DNA fragments and unincorporated radioactive dATP.  Rinse both compartments of the buffer tray with water and wipe down the broad flat vertical surface (heating plate) of the gel rig with a large kimwipe to remove any liquid buffer that has dripped down.
5)  Using a spatula labeled for radioactive-use, gently pry top plate from bottom plate only slightly at first.  This step can be very tricky, and if gel does not stick to correct plate, consult an experienced sequencer in the lab before separating plates very  much.  If gel is sticking to bottom plate, continue to separate plates and remove top plate and spacers entirely.  
6)  If using regular acrylamide gel, it may be desirable to fix the gel (though not imperative).  If you are fixing the gel, go to "Fixing Gel" instructions.  If using "long Ranger" gel, you may not fix the gel (it will dissolve).  For non-fixed gels, label a 13"x15.5" sheet of filter paper with pencil to denote any desired information about the gel, and place it directly on top of gel making sure to cover the entire gel.Press down gently to make a good contact between gel and paper.  Peel off paper with gel and go to "Drying gel" instructions.

B)  Drying Gel (30-40 min)  
1)  Place saran wrap over surface of gel.  Use two or more pieces if necessary to ensure that entire surface of gel is covered.  Saran wrap should be laid out smoothly but a few wrinkles, folds, and regions of saran wrap overlap are fine.
2)  Trim saran wrap so that no more than one inch is sticking out from the filter paper on any side.
3)  Open gel dryer, lift up rubber gasket, and place the gel, saran wrap side up, on top of the filter paper.
4)  Replace the gasket on top of the gel so that it is stretched out flat and close lid of gel dryer.
5)  Follow directions on vacuum aspirator to start vacuum.  Make sure valve to your gel dryer is open and is pulling vacuum.  Reopen lid of gel dryer and check if gasket is holding vacuum.  
6)  Press "start" button to initiate gel heating program. 
		Gel dryer program settings:
			use the first (top) program (an even drying temp.).
			temp is usually set for 80o.

C)  Removing Gel From Dryer , Checking and Exposing (15 minutes)
1)  Remove gel from dryer, turn off aspirator according to directions on aspirator, and remove saran wrap from gel or from gel dryer gasket (whichever it is stuck to).
*if saran wrap does not come off of gel easily, try to gently peel it off but if it really  is stuck, don't rip it because it will rip the gel.  You can expose the gel with the 	
	saran wrap still on if you allow for an exposure time roughly 25% longer.
2)  Check the dry gel with a geiger counter set to 1X, and record level of radioactivity on sequence data sheet.  [Note:  if the reading is greater than 4K, then reset the geiger counter to 10x to read the value.]
3)  Bring gel and data sheet into dark room.  
4)  Select a metal cassette that is facing green side out (unused) and place on counter. (see below if there are no metal cassettes available).
5)  Open cassette and place gel, gel-side-up, in cassette.
6)  Turn on red light, turn off regular light, remove film from drawer, place one piece of film on top of gel inside cassette.  Seal cassette, close up film packaging, replace film in drawer and turn on regular light.  If your gel loading configuration was symmetrical (ie it would appear similar flipped over), you should take measures to ensure that you do not lose track of the correct left-right alignment of the film (making a small mark on one side of the film is usually sufficient).
7)  Put a small piece of tape on cassette labeled with your name and the date of exposure.
8)  Replace the cassette, red-side-out, in the rack and record which cassette your gel is in on data sheet.
*If there are no metal cassettes available, use a cardboard cassette by placing gel and film in cassette with lights out as above, sealing cassette and film package as above, then place cassette in between two glass plates (in drawer) and clamp glass plates together along all edges.
9)  Leave the cassette in the rack to expose for 1-7 days (or more if you're desperate, it is difficult to over-expose when using 35S) depending on strength of radioactive signal on gel.  A gel that reads 2K or more in step 2 (over the entire gel, not just the top) should yield readable sequence with an overnight exposure.  If the gel reads 0.5-2K, a period of roughly 3 days is usually necessary for exposure.

D)  Developing Gel (10 min. => 25 min.)
1)  Make sure fluids in both fixer and developer tanks below automatic developing machine have at least 2 inches of fluid in them (if not, these fluids need to be replaced).
2)  Turn water valve (blue handle near door) into vertical position (on) and turn on automatic developer by pressing two adjacent black switches on left (west) side of machine 
3)  Wait approximately 15 minutes for machine to warm up (indicated by red light flashing on and off above black switches).
4)  Feed one piece of old film (sitting on top of machine) through feed tray on right (east) side of machine.
5)  Place cassettes with films to be developed on counter and turn off lights (except for red light).
6)  Open cassette and label the top left-hand corner with your intials and the gel number (e.g. TCG 94-15).
7)  Take the film to the X-ray processor and feed the film into machine through feed tray.  The rollers can be activated by pressing the black button next to the feed tray or by gently pushing the film into the processor.
8)  When the machine buzzes, you may feed in a new film or turn on the lights again.
8)  After three or four minutes, the machine will start spitting out the films in the order that you put them in.  Remove films and label them with the order of samples in lanes, primers used, length of exposure, length of gel run,and other relevant information (date, experimental treatments, etc.).

References

Aguadé, M. W. Meyers, A. D. Long, and C. H. Langley. [1994?]  Reduced DNA sequence polymorphism in the su(s) and su(wa) regions of the Drosophilia melanogaster as revealed by SSCP and stratified DNA sequencing.  I believe that is now published in PNAS.
Bruford, M. W. and R. K. Wayne. 1993.  Microsatellites and their application to population genetic studies.  Current Opinion in Genetics and Development 3: pages not assigned to preprint.
Clinton, M. and R. K. Scougall. 1995.  Detection and capture of 35S-Labeled gas released from reaction tubes during differential display PCR.  BioTechniques 19(5):798-799.
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Appendix I.  Fixing gel (25 minutes)
1)  Place glass plate with gel on top in the fixing tub (gel side up).
2)  Gently pour in 1 liter of fixing solution (5% acetic acid, 5% methanol) so as to cover the gel completely.  Do not disturb gel by pouring in fixing solution too fast.  Folds, bubbles, and floating portions of the gel may be eliminated by gently touching the gel while in the fixing solution.  
3)  Allow gel to sit 15 minutes in fixer.
4)  Cut out a 13"x15.5" piece of filter paper, write any desired information about the gel in pencil on the filter paper, and place it in the fixing tub directly covering the entire gel.  
5)  Remove fixing solution with aspirator hooked up to faucet while holding filter paper in place over gel with other hand.
6)  Lift plate out of fixing tub and place, gel-side-up, on a flat counter surface with bench paper.  
7)  Cover entire filter paper with small stacks of paper towels (0.5 inch stacks) to suck up moisture.  Flip over paper towel stacks, press down again gently, and remove paper towels.  
8)  Gently peel up filter paper with the gel now stuck to it.