General naming scheme (for gel images and probes, but not for genes or clones):
   Genus species cultivar Type - number . version 
     GmwPb-120.2
   
   Use a "wild-card" letter (x?) if necessary. For example, a gel might contain 
   BACs from more than one cultivar. Then the gel name would start Gsx
  
  Species and library names
    Species   library
    prefixes  name
      Gmw1     gmw1       Glycine max - Williams Hind III Library
      Gmw2     gmw2       Glycine max - Williams Bsty1 Library
      Gmp1     gmp1       Glycine max - PI996983
      Gtd1     gtd1       G. tomentella diploid "t3 species"
      Gtt1     gtt1       G. tomentella  tetraploid "t2 species"
      Mt       mth2       Medicago truncatula Hind III 2 library
      Pv       ??         Phaseolus vulgaris (collaborators)
      Tl       tl1        Teramnus labialis
  
  BAC:    
   library-plate row col; no leading zeros; - not _; lower case:
     mth2-5p10
  
  Gel images:
    Phase 1 gel im:      
      GscPhI-1.1.tiff
      
    Other gel im:      
      GscGel-1.1.tiff
  
  Contig files (FPC to species directories at IU site)
    IM3 folder:    
      GscIm3-123.2
  
    FPC gel image:
      GscFPCgel-123.1.tiff
      
  BAC end sequences    
    Genbank accession
      AW123456 
        optional: add 
          Genus species cultivar plate well column -forward/reverse . version
        AW123456-Gsc5p10-f.1 or -r.3 etc.
  
  Gene
    "Long name": For most purposes, we will use the University of Oklahoma gene 
    naming scheme (see the gene calls in the OU GMOD browser), with an extra 0 
    added to gene numbers to provide room for additional genes. For example,
      AC126014.fg.140  
    The parts of the name are: 
      Genbank accession; gene-calling program; gene order on BAC x 10
    
    "Short name": For some comparative work (e.g. contig sketches, phylogenies),
    it will also be helpful to use a shortened name that includes
    Genus, species, cultivar, and clone: 
      Gsc5p10.140 Gsc + plate 5 row p column 10 gene 14 (by gene-calling 
      method specified in a legend for the figure, e.g. Fgenesh or Genscan). 
    
    Optionally, include the the gene-calling method and other essential 
    information before the gene number, e.g. Gsc5p10.fg.1 for an FGenesh call. 
    The method for establishing short names for a BAC or contig will need to
    be briefly described in a figure or phylogeny legend.
    
    Distinguish occasional duplicate BACs per well with a letter following
    the clone address e.g. Gsc5p10a.140 . 
    
    For genes identified before before complete BAC sequencing (e.g. from an
    end sequence), Genes should simply be called in order of naming: 
    Gsc-gn1.1, Gcs-gn2.1 etc.
    
    How to track these names: the accession-based names will be visible in the
    OU GMOD browser. We will also  download the gene names and predicted ORFs 
    from OU GMOD browser and enter them into our database. 
  
  Probe
    GscPb-123.1
  
  Clone
    pGmcPb-123.1
  
  Primer:    Pm-123.1
    Primers may not be species-specific, so are just indicated as 
    "Pm" for "primer".

PREHYBRIDIZATION

1. Preheat Hybridization Buffer to 60oC in a water bath (this usually takes
between 10-30 minutes depending on how much hybridization buffer is in the
container.  If you are in a hurry, you can microwave the solution for not
more than 1min at a time.  Then place the container in the 60oC water bath.
 Do not overheat the solution).

2. Turn on the following heating equipment:
 a. Hot plate/Stirrer - Highest setting
 b. Heating block - 105o C
 c. Hybridization Incubator - 60oC

3. Boil frozen salmon testes (st)DNA for 15 min in a beaker of water on the
hot plate.

4. Prepare hybridization container (Rubbermaid 10 x 13 in. with cover).

5. Five minutes before the stDNA has completed boiling add 200-300ml of
hybridization buffer to each hybridization container.  Volume of
hybribization buffer dependent on the number of filters hybridizing; 200 ml
for 1, 300 ml for 6 filters; filters should be covered with solution.

6. Add 3.6 ml of boiled stDNA for 200 ml of buffer and 5.4 ml stDNA for 300
ml.

7. After stDNA has been added, mix the solution evenly by gentle swirling
two or three times.

8. Add the filter(s) one at a time to boxes, DNA side down.  If putting
more than one filter per box, make sure that the first filter is completely
wet before adding the next filter. 

9. Stack the containers (if using more than one container). Use container
lid as cover for top of stack.  Stack the containers carefully to prevent
evaporation of hybridization buffer. Place a 1L beaker of water in
incubator to slow evaporation of hybridization solution.

10. Pre-hybridize the filters for at least 3.0 hours at 60oC.


RANDOM HEXAMER LABELING
  
1. Start the labeling step approximately 1.5 hours after the filters have
been placed at 60oC for pre-hybridization.

2. Thaw the 32P. Take the  dNTPs, reaction buffer, and  distilled water out
of the freezer (Invitrogen Random Primers DNA Labeling System) and place on
ice.  Thaw the probe template DNAs. Label 0.5µL microfuge tubes (these do
not have to be sterile) for the labeling reactions.

[NOTE:  when hybridizing  more then 2 filters in a single container label 2
tubes of probe for the hybridization]

3. Calculate the volume of probe template DNA that contains approximately
25-50ng.  

4. Calculate the volume of H2O to add to the DNA, 32P, dNTPs, reaction
buffer, and Klenow for a total volume of 50µl. 

5. In each labeled tube, first add sterile double distilled H20, then ~40ng
template DNA.  Boil for 8 min in a heating block or water bath.  Cool on
ice.

6. Once cool, add, on ice, IN ORDER:

6 µL dNTPs
15 µL reaction buffer
1 µL Klenow

Note: make sure the 32P is thawed before adding the Klenow.  If not, store
the tubes on ice until the 32P is ready.  

7. Add 5 µL 32P to each tube. Vortex the tubes slightly.

8. Incubate the tubes in a heating block at 37oC for between 60 and 90
minutes.



STOPPING THE LABELING REACTION AND STARTING HYBRIDIZATION

1. Remove the labeling tube from the heating block.

2. Add 190µL blue stop solution.  

3. Transfer the tube to the 100oC heating block and boil probe for 8min
with the lid open. 

4. Take the hybridization container from the 60oC incubator. Remove the lid
and place under the hybridization container.

5. Remove the probe from the hot block.

6. Lift one edge of the filter(s) and pick it up with your left-hand (if
there is more than one filter per box, lift the filters together on one
edge).  Let the filter(s) drip inside the box.

7. While the filters are out of the box, pick up the corresponding tube of
probe solution.  Hold the tube by the side and pour the probe into the
hybridization box.  Mix evenly using a gentle swirling motion.  

8. Replace the filters into the hybridization box, one at a time, making
sure that each filter is covered with solution.  Do not let the filters
move up to the sides of the box.

9. Repeat for each hybridization container.  Stack each container on top of
the other.  Replace the container lid on top of the stack.

10. Hybridize overnight at 60oC (24 hrs is desirable).


WASHING 

1. Prepare SSC of the appropriate stringency for your first wash (2X SSC
and 0.1% SDS: 100ml 20X SSC plus 5ml 20% SDS per liter of wash solution). 
Use approximately 1000 ml of wash solution per container.  Pre-heat the
wash solution to 60oC in a waterbath.

2. While waiting for the wash solution to heat up prepare the appropriate
number of wash containers and screens; wash up to 8 BAC library filters per
container.  Place one screen at the bottom of each container and a screen
between each filter.

3. Pour the appropriate amount of wash solution into each wash container. 
 
4. Using forceps lift one filter, holding the edge with your left hand, and
let the filter drip into the hybridization container, while with your right
hand reach for a screen.

5. Place filter into a wash box with forceps and cover with screen.  Push
down the screen gently with the tip of a forceps until it is covered
entirely by the wash solution.

6. Repeat with the remaining filters.

7. Cover each wash container tightly.  Shake the filters (55rpm) at 60oC
for 15 minutes.  Pour off the wash solution and replace with second wash
(1X SSC and 0.1% SDS: 50 ml 20X SSC plus 5ml 20% SDS per liter of wash
solution).  Do the same with the third wash (1X SSC and 0.1% SDS). 

8. Pour off as much of the third wash solution as possible.  Lay the
library filters onto a piece of saran wrap, DNA side down.  Blot the
filters dry with paper towel, and wrap in the plastic wrap.  Monitor the
radioactivity level in each filter using a Geiger counter.

9. Take the filters to the dark room.  Do the following steps in complete
darkness or at most only with the red safety light on. 

a. Place the filters in film holders, DNA side facing up, label x-ray films
with the filter numbers (in the right hand corner), lay on filters.

b. Close the film holder, place in -80oC freezer. 

10. Exposure will be 48 to 72 hours depending on the counts.


DEVELOPING FILM

1. Take the appropriate filters out of the -80oC freezer and allow to thaw
for 5-10min.

2. Develop films.

3. Remove the filters from the film holders and put them in the
refrigerator until stripping.  Do not let your filters dry out before
stripping.


STRIPPING FILTERS


1.  Peel the Saran wrap from the filters.

2. Place the filters, without screens, into 1 L (for up to 20 filters) of
0.05N NaOH.  Shake for 15min.  

3. Pour off the solution and add 1L (per 20 filters) 0.2M Tris pH 7.5, 0.1X
SSC, 0.1% SDS (200ml of 1M Tris pH 7.5; 5.0ml 20X SSC; 5.0ml 20% SDS).
Shake for 15min.

4. Pour off the solution and add 1L 0.1X SSC, 0.1% SDS.  Shake for 10min.

5. Reuse the stripped filters immediately or lay flat on a sheet of Whatman
3MM, DNA side up and cover with another sheet of Whatman.

6. Air dry overnight.  

****************************************************************

PLEASE OBSERVE RADIATION SAFETY RULES AT ALL TIMES DURING THIS PROCEDURE! 
WEAR LABCOAT AND GLOVES AND MONITOR YOURSELF AND THE 32P AREA CAREFULLY!!

How to identify an address:

The membrane is divided into 6 fields (diagram provided).  Each field
contains 384 squares.  The 384 squares represent the row and column
identification of the BAC.  Within each square there are 16 positions where
8 clones are spotted in duplicate (diagram).  The pattern of the spotted
clones will generate the plate address of the BAC.  To identify your clone,
please follow the directions below.

The most complicated part about identifying a clone address is that
consecutive plates are not spotted into each field.  The 384 well plates
are spotted onto the membrane with plates 1-6 spotted into fields 1-6
respectively (duplication pattern 1, see diagram).  Since there is a total
of 6 fields on the membrane, the cycle will continue with the next six
consecutive plates (plates 7 through 12) again being spotted into fields 1
through 6 respectively, but in a different duplication pattern (duplication
pattern 2, see diagram).  This gridding cycle will continue until all the
plates have been spotted.


1. The library name and filter number is used to orient the membrane. 
Place the membrane with the label facing up and on the right-hand side as
shown in the diagram. (The colonies are on the same side of the filter as
the label)
2. Identify the field number of the hybridizing colonies.  The spacing of
colonies is slightly wider between the fields.
3. Identify the well location (I have included a grid to help locate well
positions) and identify the well position (e.g. L18)
4. Identify the plate number.  This is accomplished by determining the
orientation of the duplicate spots (duplication pattern in the diagram) and
referring to the table inside each field in the figure.
5. Libraries that have more than one filter (48 plates) will also need to
decode the plate number based on the filter.  Plates 1-48 are spotted on
filter A, 49-96 on filter B, etc.  Identify the plate number and well
location as described above and record the filter letter.  Go to the
conversion table and read down the column corresponding to the filter
letter and read across the plate number identified from the filter.  The
intersection is the actual plate number of the clone.


Example:  If you have horizontal spots, they could either be duplication
position 4 or 8 (from the duplication pattern).  They are distinguished by
the closeness of the spots and position in the pattern.  Assume it is
position 4 in field 3.  Read down the table in field 3 of the diagram to
pos4 and read the plate number as 21.  If you are reading filter B,
identify the library plate number from row 21 and the column labeled Filter
B of the library filter plate decoder.  The library plate number is 69. 
Once the plate number is determined, identify the well location either by
using the supplied grid or counting the rows and columns.

Library Plate Decoder

Polyfiltronics BAC Minipreps

(based on an alternative method described in the paper "Highthroughput
Fingerprint Analysis of Large-Insert Clones" M. Marra, et. al.1997. Genome
Research,. 7:1072-1084.)

Pre-culture: (Day 1 and 2) 

1.    Streak out glycerol stocks on CM/IPTG/X-gal plates: scrape frozen
culture with a sterile pipette tip, touch plate, and streak with loop.
Incubate O/N to 24 hours at 37¡C to obtain single colonies.

2. Fill a sterile Sarstedt 96-well microtiter plate [round bottom with lid
#82.1582.001] with 150 µl LB containing 12.5 µg/ml CM.  Use a toothpick to
touch and inoculate a single colony into each well.  Grow 16 hours (16-18
hours is fine) in 37¡C non-shaking incubator.

Preparation of 24-hour culture and glycerol stocks (Day 3)

3. Cover the pre-culture plate with Qiagen AirPore tape [#19571] plus
Qiagen plastic tape [#19570] on top. Vortex gently. Remove tape very
carefully to prevent cross-contamination.

4.    Fill each well of two sterile 96 well Polyfiltronics (2 ml-deep well)
blocks [from Whatman, round bottom #7701-5200] with 1.3 ml 2X YT containing
12.5 µg/ml CM.  Replicate the pre-culture into the deep well blocks by
using a flamed replicator.  Seal the blocks with a Qiagen AirPore tape
sheet.
  
5. Incubate the blocks on a slant adapter in a 37¡C shaking (300 rpm)
incubator for 23 hours (22-24 hours is fine).

6.    Glycerol stocks: Transfer 75 µl pre-culture to a 96 well Sarstedt plate
filled with 75 µl 30 % glycerol, and mix with pipette.  Seal plate with
Research Products sterile thermal seal tape [Biomek from Beckman #538619]
or sterile foil tape.  Store plate at -80¡C.

Prepare bacterial alkaline lysates: (Day 4)

Before beginning, put Sol'n III on ice and take out RNase [Sigma R-6513] to
thaw.

7. Take 96-well blocks out of shaker and place on ice for 10 minutes.

8. Pellet the cells by centrifuging the blocks at 3600 rpm (2500 g) for 15
minutes at 4¡C.  Pour off media and bleach.  Blot blocks onto paper towels.
  
9. Re-suspend each bacterial cell pellet in 150 µl Sol'n I (GET/RNAse
:0.12mg RNAse/1 ml GET-see recipe at end of protocol).  Tape each block and
vortex to re-suspend.  Let the blocks sit at room temp for 20-30 min for
the RNase treatment.

10. Add 150 µl Sol'n II (Lysis) to each well, seal block with foil tape,
and invert gently 20 times.  Incubate at room temp. for 5 minutes (no
more-begin timing when first row is added).

11. Add 150 µl chilled Sol'n III (3 M KOAc, pH 5.5) to each well, seal with
new foil tape, mix immediately by inverting 10 times, and incubate on ice
for 10 minutes.

Clear the lysates: (Day 4)

12. Centrifuge the block at 4000 rpm (3046 g) for 15-20 minutes at 4¡C. 

13. Transfer lysate (both spin and vacuum method work fine)

Spin method: Securely tape Polyfiltronics filter [from Whatman #7700-1808]
on top of a new autoclaved Polyfiltronics block so the wells line-up
perfectly. Transfer the supernatant to the filter (both Polyfiltronics
blocks will be filtered through one filter). Spin @ 4000 rpm for 35 min  at
4¡C or until the lysates are completely transferred.

Vacuum method: Place Polyfiltronics block in Polyfiltronics vacuum
manifold, place manifold cover on top, and then place filter on top-it
should fit perfectly. Transfer the supernatant to the filter. Cover filter
with plastic tape.  Apply vacuum (2-3 psi) until lysates are completely
transferred.

Ethanol precipitation: (Day 4)--make sure centrifuge is set at room
temperature

14.Remove filter and add 330 µl 100% ethanol to the solution in the blocks.
 Foil tape [3M 4 inches electrical tape] the block and mix by inverting 3
times.
  
15. Let block sit for 1 hour (RT).  Spin block at 4000 rpm for 1 hour (RT).
Do not spin for longer than an hour - you might get more DNA, but also more
junk that will give you noisy sequences!

16. Decant ethanol into waste container.  70% EtOH wash: Add 500 µl 70 %
ethanol, vortex block gently, spin for 5-10 min., pour off ethanol into
waste container, blot block on paper towels; repeat ethanol wash.  After
2nd ethanol wash, remove as much EtOH as possible.

17. Dry pellet for ~ 1 hour in vacuum. Make sure no ethanol is present!!!

18. Re-suspend pellet in 30 µl 10 mM Tris, pH 8.5 or filtered dH20 and
cover block with foil tape and wrap with saran wrapà suspend O/N (4¡C)  To
sequence, use 8 µl of DNA with 2 µl of 5 pmol primer (full reaction).

****BACs can also be suspended in 15 µl of dH20 for sequencing.  Then use 4
µl of DNA with 2 µl of 5 pmol primer to run half-reactions (don't adjust
primer in half reactions).

Day 5: Vortex block, spin down briefly, transfer DNA to autoclaved MJR 96
well PCR microtiter plate, and cover plate with tape for long-term storage.
 Store at -20 degrees Celsius.



Solution I:  GET/RNAse (100 ml)
                                        FINAL
2.5 ml   2.0 M Glucose (filter sterilized)         50 mM
2.5 ml   1.0 M Tris, pH 8.0                    25 mM
2.0 ml   0.5 M EDTA, pH 8.0                    10 mM
dH20 to 100 ml

Autoclave 15 minutes.

Add RNAse just before use: 

    Add 20 µl RNAse (10 mg/ml stock) per 1 ml GET 
Final conc: 0.12 mg RNAse/1 ml GET

Solution II: Lysis buffer (0.2 N NaOH/1%SDS)

Make just before use.

Add one volume 2% SDS to one volume 0.4 N NaOH.

Mix gently.

Solution III: 3 M Potassium/5 M Acetate, pH 5.5 (100 ml)

29.4 g Potassium acetate
28.5 ml glacial acetic acid (start with 25 ml and check pH as more is
added)
25 ml ddH20 

Adjust pH to 5.5 and bring volume to 100 ml with ddH20.

Note:  the pH can be 5.3-5.5

Autoclave 15 minutes.

1.  Digest BAC DNA with EcoRI or HinDIII depending on the BAC library

a. Digest 1 µl of BAC DNA with 2 units enzyme in a total digest volume of
   10µl.
b. Incubate @37degrees for 3 hours. Add 4 ul 1:1 "Blue Juice".
c. Heat @65 degrees for 5 min. Load 7 ul on gel.


2. Load digested BAC DNA and molecular marker on a 1% agarose gel.

a. Use a 20 x 25 cm gel tray with two 30 or 42 tooth 1.5mm combs (depending
on
   the number of positive BACs).
b. Load 5µl Marra marker, skip a lane, and then load 5 lanes with 7 µl
digested
   BAC DNA, repeat.
c. Run gels in 1X TAE for 17 hours at 26 volts (DNA will have migrated to
   middle of gel tray)

3. Southern Blot gel

4. Hybridize Blot with same probe that pulled out BACs and include labeled
HinDIII ladder to visualize Marra Marker

5. Expose to film

6. Develop film

7. Using a ruler (if necessary), group positive BACs into preliminary
contigs based on similar hybridization patterns. Mark each group with a
different color if there are several present.  You can put BACs together
that you feel uncertain about---simply place a question mark behind them so
that they are carefully examined after PhaseII gels are run.

8. BACs that do not light up are recorded as false positives and not
included in Phase II gel.

9. Run Phase II gel (fingerprint gels)

Process gel for blotting

1. Run and stain gel, as usual (there are usually two portions for every
gel, top and  bottom. Carefully cut the gel into top and bottom portions,
cut off the bottom 1 cm of each portion, clip off corners to note
orientation, and process both portions in parallel).

2. Soak gel in 600 to 800 ml of 0.25 N HCl for 10 minutes (exactly) on
shaker. Blue dye should change to yellow color during this step. This step
depurinates the DNA.

3. Pour off HCl and add 800 to 1000 ml of 0.5 N NaOH/0.5 M NaCl. Place on
shaker for 30-45 minutes. Dye should change back to blue during this step.
This step denatures the DNA.

4. While gel is in NaOH/NaCl, prepare membranes for blotting.

a.For each gel to be blotted (assuming upper and lower portions), prepare:
6 pieces of Whatman 3MM, 21 x 12 cm in size, and 2 pieces of Hybond N+, 20
x 10 cm in size. Handle membranes with gloves.

b.    Use permanent ball point pen to write number of the blot at the bottom
of the Hybond membrane (write on the side in which the middle curves
upward).

c.    Soak Hybond membrane in distilled water for five minutes.

d.    Pour water off membrane and add 0.5 N NaOH/0.5 M NaCl. Soak in this
solution for 5 to 10 minutes.

Set up Southern Blot

1. Set up blotting apparatus by adding up to 2 L of 0.5 N NaOH/0.5 M NaCl
and squeezing bubbles out of the sponges.

2.    Briefly soak two Whatman 3MM filters in 0.5 N NaOH/0.5 M NaCl and place
on top of one of the sponges. Repeat this process by placing two soaked
Whatman filters on top of the other sponge. Use pipet to roll out any
bubbles.

3.    Carefully place the upper portion of the gel on upper sponge. Place
lower portion of gel on lower sponge.

4. Surround gels on all sides with cut strips of thin plastic sheets. Make
sure that no surface of the sponges shows through.

5. Carefully place the numbered Hybond membranes over top and bottom
portions of gel with numbers facing up.

6. Briefly soak one piece of Whatman in 0.5 N NaOH/0.5 M NaCl and place on
top of upper portion of gel. Repeat with lower portion of gel. Use pipet to
roll out any bubbles.

7. Place one half of a stack of paper towels carefully over each Whatman
filter.

8. Place plexiglas sheet and a tray (with just enough water to cover its
bottom) on top of the paper towels.

9. Allow blotting to continue for 6 hours to overnight.

Process Southern Blot

1. Remove paper towels, and place Hybond membranes in 0.4 N NaOH for one
minute (you may also wish to stain the gel to make sure that all DNA has
been  transferred).

2. Pour off NaOH and neutralize membrane(s) by adding 0.2 M Tris (pH
7.5)/2X-SSC.

3. Place membrane (DNA side up) on Whatman filters and dry for several
minutes.

4. Transfer membranes to new Whatman filters and bake in vacuum oven at
70¡C for 2 to 3 hours.

5. Store blots at room temperature.

PREHYBRIDIZATION

As for library hybridization except use the following volumes of
hybridization buffer:

filters        Hyb. buffer        stDNA
1             50mL                0.9mL
2             + 15mL            + 0.25mL
3            + 15mL            + 0.25mL
4            + 10mL            + 0.20mL

RANDOM HEXAMER LABELING  

As described for the library hybridizations.

STOPPING THE REACTION AND STARTING HYBRIDIZATION

As described for the library hybridizations.

WASHING 

As described for library hybridization except:

Use approximately 100ml of wash solution per filter.

EXPOSING FILM

Exposure times:

a) 150-300 cpm - 3 days
b) More than 300 cpm - 1 to 2 days

DEVELOPING FILM

As described for library hybridizations.

STRIPPING FILTERS

As described for library hybridizations.

1. BAC digests for fingerprinting gels (aim for ~50 ng/lane)

Make a 10 ul digest:

DNA                    7 ul            
dH20                    0.4 ul
10X BSA                    1.0 ul
RI Buffer                1.0 ul
40mM spermidine (optional)    0.5 ul
EcoR I (NEB)                0.1 ul

Mix and digest for 3 hours @37¡C.  Add 2 ul conc. Blue Juice (bromophenol
blue + glycerol).  Load 8 ul or as much as you can on the gel.  Load 10 ul
Marra Marker after every six lanes of digests (load an even number of BAC
lanes in between the markers).

2. Start preparing gels at least two hours before you plan to load them.

3. Prepare 2.5 L of 1X TAE buffer for two gels (enough to make gels and
fill gel rigs):

a. Measure out buffer for your gels
b. Cool remaining buffer:
- Make buffer the day before and store @ 4¡C
- or make several hours ahead of time and store in the -20¡C (-80¡C if you 
are really in a hurry) until the buffer temperature cools to 8- 10¡C.  By
the time you get the gel set-up, the buffer will warm-up to the appropriate
running temperature (12-14¡C).
 
4. Make sure the tubing loops are submerged in the cooling tank, then turn
on the chiller.  Set the chiller at 2-4¡C.  It takes an hour or so to get
the chiller to the right temp.

5. Make your gels while you are waiting for the chiller and the buffer to
cool.  Preapre 1% agarose (SeaKem LE), 160 ml gels (3mm thick if gel is
20X25cm).  Use 1X TAE to make the gels.  Stir the agarose and buffer to
ensure the agarose is hydrated and evenly distributed.  Remove stir bar. 
Weigh beaker containing agarose solution.  Add a couple grams of water. 
Cover mouth of beaker to reduce evaporation.  Microwave until boiling
(about 5 minutes).  Take beaker out at 1:30min intervals and swirl to
ensure even mixing of agarose.  Take beaker and weigh.   Add water to bring
beaker to original weight.  Swirl beaker and microwave for another 30
seconds.  Weigh one last time to ensure beaker is at its original weight. 
Place beaker in 60¡C water bath until cooled to about 60¡C, usually about
ten minutes.

6. Tape gel tray and warm the tray for about ten minutes in  55-65¡C oven. 
Pour gel and put one 1 mm 42 tooth comb in place.  Let solidify for at
least 30 minutes before using.  If you have time, cool the gel in the
fridge.
 
7. Put together the re-circulating rig.  Intake is at the top of the gel
(comb end), outflow is at the bottom.

8. Make sure the gel rig is level.  Place a bubble level in the middle of
the rig.  Place paper towels under the rig to level.
  
9. When the buffer has cooled to about 10¡C, add it to the gel rig.  Turn
the pump up to about 90 to get the bubbles out of the tubing.  Then set the
pump at 40 to get buffer to running temperature (make sure you do this at
least 30 minutes before you load your gel).

10. Remove the tape from the ends of the gel tray. Turn off the pump for a
few minutes and place the gel tray in the gel box.

11. Make sure (use a ruler) that there is no more than 3-5 mm of buffer
covering the gel.

12. If you have time, pre-run the gel for 15 minutes at 72V with pumps
running.  Check the buffer temperature; it should be about 12-14¡C.  While
you are waiting, heat your gel samples and markers in the 65¡C water bath
for 5 minutes.  Spin down the samples and place on ice.

13. Turn off the pumps to load the gel.  Take the temp. of the buffer. 
Load the gel and set the voltage to 90 for 5-10 minutes so that the samples
can run into the gel.  Set the voltage back to 72V and turn the pump back
to 40.  Record the time.  Run gel at 72 V for 17 hours.  Check the final
amps and temp.
 
14. Thaw 40 ul of Sybr Gold (keep it in the dark).  You will want to make a
1:10,000 dilution of Sybr Gold (1ul Sybr Gold/10 ml buffer).  Measure out
400 ml of 1X TAE and pour into a foil wrapped 7.8 L Rubbermaid box w/cover.
 Place gel on a thin piece of plexiglass and place in the box.  Add 40 ul
of Sybr Gold to the buffer.  Cover the box and check that the foil covers
the box well.  Stain gel for 1 hour by shaking moderately fast, being
careful not to tear the gel.  It is not necessary to de-stain the gel.
 
15. If you have a good gel documentation system like Bio-Rad's gel
documentation system take a picture and save as TIFF file.

***Suggestions***

- For FPC it is important to set up the gel so lane spacing remains
constant over the whole gel.  Our gels are loaded with 1 lane Marra marker
,5 lanes BAC DNA, 1 lane Marra marker, 5 lanes BAC DNA, etc. 

- Drill 3 small holes (4mm) equally spaced at top and bottom of gel tray,
to keep the gel from floating (remember to tape holes before pouring gel).

- The temp. of the gel rig should be 12-14¡C (aim for 13¡C, no more than
15¡C)
 
- Set the pumps between 50 and 60
 
- Experiment with voltage and time:
    - Try 90-95 V for 20 hours
    - Try lowering the voltage if you don't want to run the gel as far

- 56 V for 16 hours also seems to work well (run at 14¡C so it runs a
little further).  Or try a voltage somewhere in between 56 and 95 V.
Pay careful attention to changes in temperature and quality of the gel.

Marra Marker 

The following molecular marker is used for both Phase I (DNA is transferred
to nylon and hybridized) and Phase II gels (DNA is cut with EcoR I and run
on a long gel for FPC analysis).  The marker recipe is based on methods
used by M. Marra, et al. (High Throughput Fingerprint Analysis of
Large-Insert Clones).

Note: Marker III is lambda cut with HinD III and EcoR I (0.12 - 21.2 kb
fragments)

Concentrated Marra Marker (do not load this)

                                    Stock            Volume to add
1 kb Ladder (Life Technologies)            1 µg/µl             8.30 µl
( cut with HinDIII (Life Technologies)        500 ng/µl         16.65 µl    
Marker III (Boehringer -Mannheim)            250 ng/µl         33.3 µl
TE (10:1), pH 8.0 (10 mM Tris, 1 mM EDTA)                          941.75 µl
Concentrated Blue Juice (5X)                                 250.00 µl
                                                    1250.00 µl total 

Mix well.  Enough to make ~2400 µl Ready to load Marra Marker.  May be
stored at -20 ¡C.

Ready to load  Marra Marker: 400 ul aliquots (10 ng/(l)

                                    Volume
Concentrated Marra Marker:                200 µl
TE (10:1), pH 8.0 (10 mM Tris, 1 mM EDTA)        170 µl
Concentrated Blue Juice (5X)                30 µl
                                    400 µl total

Mix well.  Store at 4¡C.
  
Before loading, incubate in 65¡C water bath for 5 minutes.  Put on ice. 
Load 7 µl on a Phase I (Load 1-2 µl next to genomic lanes) or 10 ul on a
Phase II gel. 


Marra Marker Molecular Weights:

1 kb (BRL), ( cut with HinDIII (BRL), Boehringer Mannheim Marker III (( cut
with EcoRI and HinDIII)

23,103                    bold=( fragments
21,226
12,216
11,198
10,180
9416
9162
8144
7126
6557
6108
5148
5090
4973
4361
4268
4072
3530
3054
2322
2036
2027
1904
1636
1584
1375
1018
947
831
564
506
396
344
298
220
201
154
134
125

    

Disclaimer:
Any opinions, findings, and conclusions or recommendations expressed in this material are those of the author(s) and do not necessarily reflect the views of the National Science Foundation