Model Organisms

Verifying Genomic Integration

Lysing cells and setting up reactions

  • Single colony PCR is used to identify transformants which have the gene of interest integrated at the genomic locus. One set of PCR primers is designed to identify whether the prototrophic or auxotrophic allele is at the genomic locus. In this case, one primer binds to the genomic DNA and the second primer binds to the coding sequence of the auxotrophic gene. The second set of PCR primers is designed to identify whether the Sikorski vector is at the genomic locus. In this case, one primer binds to the genomic DNA and the second primer binds to the Sikorski vector.
  • Typically, 3 transformants are screened per desired strain. Re-streak each colony onto the appropriate selection plate.


  • Pick a colony at least 2 mm in diameter using a wooden toothpick. Resuspend it into 11 uL Lyse-N-Go (Pierce) in a thick-walled PCR tube. Transfer 5.5 uL of the mixture into a second PCR tube.
  • Load all the tubes into the PCR machine and run the Lyse N Go PCR protocol.
    • 65 C for 0.5 min
    • 8 C for 0.5 min
    • 65 C for 1.5 min
    • 97 C for 3 min
    • 8 C for 1 min
    • 65 C for 3 min
    • 97 C for 1 min
    • 65 C for 1 min
    • 80 C forever
  • DO NOT remove the tubes from the pcr block for the next step.
  • Add 45 uL PCR mix to each tube, while in the pcr block. Pipet up and down to mix. For each colony, you will have two PCR reactions corresponding to the two sets of PCR primers. The PCR mix is composed of:
    • 5 uL 10x Thermopol buffer
    • 1 uL 10 mM dNTPs
    • 0.25 uL 100 uM forward primer
    • 0.25 uL 100 uM reverse primer
    • 0.5 uL NEB Vent polymerase
    • 38 uL water (to 45 uL total)


  • Alternatively you can use the zymolyase/epitaq protocol which usually works better
  • Pick a colony at least 2 mm in diameter using a wooden toothpick. Resuspend it into 15 uL Zymolyase in a thick-walled PCR tube.
  • Load all the tubes into the PCR machine and run the Zymo protocol.
    • 37 for 15 minutes
    • 95 for 5 minutes
    • 10 forever
  • The PCR mix is composed of:
    • 5 uL 1:5 dilution of lysed cell mix
    • 5 uL 10x Eppendorf buffer A
    • 1 uL 10 mM dNTPs
    • 0.5 uL 100 uM forward primer
    • 0.5 uL 100 uM reverse primer
    • 0.25 uL NEB EpiTaq polymerase
    • 37.75 uL water (to 50 uL total)

Zymolyase solution

  • Make 2.5mg/ml zymolyase solution in 0.1M Sodium Phosphate buffer pH 7.5
  • Mix by inverting, not all will go into solution
  • Spin down and aliquot supernatant for storage at -20C


  • Primer sets for each locus:
Sikorski vector to be integrated allele primers (positive) vector primers (negative)
pRS304*(TRP1) DL11, DL17 DL11, DL22
pRS305 (LEU2) DL28, DL29 DL28, DL30
pRS306 (URA3) IP108, IP111 IP108, DL18
  • Run the following PCR cycle:
    • 95C for 2 min
    • 95C for 30s
    • 53C for 30s
    • 72C for (URA = 1:00; TRP = 1:30, LEU = 2:00)
    • repeat 35 times
    • 8C forever
  • Run the PCR products on an agarose gel. Since the bands are typically rather faint, pour the gel as thick as possible, and load 35 uL PCR + dye in the 5 mm-wide lanes.
  • Use the following table to determine whether the integration was successful.
gene at locus pcr product size
allele primers vector primers
TRP1 435 1031
trpD63 none none
LEU2 1168 1464
leu2D1 none 979
URA3 670 none
ura3-52 699 651

Primer Sequences


Archiving Yeast Strains

  • Inoculate YEP media with a single yeast colony.
  • Grow overnight at 30C in a roller drum.
  • Mix 50% glycerol and yeast culture in a 2 mL screw-top vial to give a final glycerol concentration of 15%(v/v), e.g. 300 uL 50% glycerol and 700 uL yeast culture.

Yeast Transformation Cell Competency And Transformation Protocol Schiestl Gietz

  1. Pipette 5 ml of dropout media into a sterile 17 x 100 mm, 14 ml polypropylene, round bottom tube (e.g. Falcon). Inoculate media with yeast from a growing colony. Shake in 30??C incubator at 250 rpm overnight. You may want to inoculate several tubes — use the one which grows fastest for the remainder of the procedure.
  2. Vortex the yeast tube (to disaggregate HF7c yeast strains) and add the contents to a sterile 125 ml flask containing 45 ml of YPD. Shake the flask containing the 50 ml mixture in a 30??C incubator at 250 rpm for 5-12 hrs (optimal incubation time is empirically-derived).
  3. Pour the contents of the flask into a sterile 50 ml Corning centrifuge tube and place the tube into a fixed-angle rotor within centrifuge. Balance tube and spin at 7000 rpm for 5 min. Pour off supernatant.
  4. Resuspend pellet in 50 ml of ddH2O by vortexing. Repeat centrifugation at 7000 rpm for 5 min.
  5. To aspirate all of supernatant without disturbing pellet: discard supernatant by inverting 50 ml tube and keeping it inverted, pick up a micropipetter with the volume set to 1000??l. Immediately after positioning the 50 ml tube upright, suck out any residual liquid that gravitates to the tube bottom (usually around a few hundred??l).
  6. Resuspend pellet in 1 ml of 100mM LiAc and transfer resuspension to a microfuge tube.
  7. Spin microfuge tube on high for 20 sec and aspirate supernatant.
  8. Add a volume of 100mM LiAc to pellet to bring solution to a final volume of 0.5 ml (usually around 400??l of 100mM LiAc).
  9. Thoroughly resuspend pellet by vortexing and/or micropipetting up and down.
  10. Prepare a 50??l aliquot in a microfuge tube for each transformation desired.
  11. Spin microfuge tube on high for 20 sec and carefully aspirate supernatant.
  12. Add transformation indgredients to pelleted yeast in this order:??
    1. 240??l of 50% PEG (buffers cells from 1M LiAc)
    2. 36??l of 1M LiAc
    3. 25??l of Salmon Sperm DNA (I incubate this at 95??C for five minutes before using)
    4. 50??l of plasmid/ddH2O solution (I use 50??l of plasmid when library screening)

  13. Vortex tube ~30 sec to resuspend pellet in transformation mixture.
  14. Incubate tube 30 min at 30??C.
  15. Heat shock tube 20-25 min at 42??C.
  16. Spin microfuge tube at 8000 rpm and carefully aspirate supernatant (don’t want to lose any transformed yeast).
  17. Very gently resuspend pellet in 1 ml of ddH2O by micropipetting up and down.
  18. Plate on dropout media. Spread just enough to expedite uptake of liquid by the media. If plating on 150 mm plates, spread 250??l of transformed yeast suspension.
    If plating on 100 mm plates, spread 100??l of suspension along with 100??l of ddH2O.
  19. Incubate plates at 30??C for 3-5 days and look for colonies.

Competent Cells


  1. Pick one colony off fresh DH5() plate into 2.5 ml LB supplemented with 25??l 1M MgSO4??(10mM final conc.)
  2. Shake at 37??C overnight and until use
  3. Do a 1:500 dilution by inoculating 100 ml of SOB in 1 L flask with 200??l of o/n DH5(), record start time
  4. Spec starting at 3 hours after start time to an OD550??0.15 to 0.3 or “eye spec” every 20 minutes starting at 3 hours after start time.
  5. Collect in two pre-cooled 50 ml orange cap c/f tubes and incubate on ice 15 min
  6. Pellet in the hermle swinging bucket rotor at 2500 rpm for 5 min at 4??C with no brake, drain pellet thoroughly
  7. Resuspend in RF1 in 1/3 original volume(30 ml) by gently pipetting with DNA tips
  8. Pellet in the hermle swinging bucket rotor at 2500 rpm for 5 min at 4??C with no brake, drain pellet thoroughly
  9. Resuspend by gently swirling in RF2 in 1/12.5 original volume (8 ml) and incubate on ice 15 min
  10. Aliquot into pre-cooled tubes, 20 tubes with 200??l and 10 tubes with 400??l
  11. Quick freeze in liquid N2, using a different colored sharpie than the previous batch mark (see detail table this section) 200??l tubes with one slash and 400??l tubes with two slashes
  12. Store at -70??C

Using frozen competent cells

  1. Thaw in air at room temperature until just liquid, use 200??l per transformation
  2. Add DNA up to 20??l and swirl to mix
  3. Incubate on ice 30 minutes
  4. Heat shock at 42??C for 90 sec, then place on ice briefly
  5. Add 800??l LB and incubate at 37??C for 45 minutes
  6. Plate out under appropriate conditions


  • Once iced, keep cells cool and pre-cool all tubes
  • Be gentle, don’t vortex or roughly pipet cells
  • Use forceps to lower tubes into liquid N2??in case tubes leak

Sequencing Procedure

Plate Preparation

  1. If new plates, mark the treated side of the plate with an RVS using the diamond pencil.
  2. Using Alconox soap and glass sponge, clean larger (non-treated) plate with small circular motions including corners in dish sink using styrofoam box for support.
  3. EtOH and wipe clean with kimwipe.
  4. Lay on two green test tube racks and EtOH/kimwipe dust off.
  5. Place side spacers with one foam end.
  6. Repeat cleaning as in step 1 & 2 on treated glass (smaller plate).
  7. Treated plate can be used 3-5X between treatings with Rain-X(see below).
  8. Check treated plate for dust and clean.
  9. Place plate treated side down onto of spacers.
  10. Position bottom spacers so seal if formed.
  11. Place one clip on each side and two clips on each corner, 5 clips on each side total.

Rain-X Plate Treatment

  1. Clean smaller treated plate as usually
  2. Squirt a small amount of Rain-X onto kimwipes.
  3. Apply to previously treated side of plate in circular motion until sol’n becomes cloudy (opaque).
  4. Repeat steps 2 and 3.
  5. Clean plate again and proceed with Plate Prep (above).

Gel mix preparation??(for 80 ml FV using Glycerol Tol. Buffer [using TBE])

  1. In 250 ml flask mix 38.4 g urea and 30 ml (26 ml) ddH2O.
  2. Dissolve by putting flask in pan H2O bath on stir plate with heat on #3.
  3. Add 1/2 to 1 g of Mixed Bed Resin.
  4. Add 12 ml 40% 19:1 acrylamide/bisacrylamide.
  5. Stir till dissolved, do not leave unattended.
  6. While urea is dissolving, filter 4 ml 20X Glycerol Tol. Buffer (8 ml 10X TBE). IF using TBE for buffer, first filter 70 ml 10x TBE buffer to make buffer chamber solUn. Leave 4 ml (8 ml) in bottom of filter unit.
  7. Make 10% APS by dissolving 0.1 g into 1 ml of ddH2O in 1.5 ml bullet.
  8. Once the urea sol’n is dissolved, filter it onto the 4 ml (8 ml) of filtered buffer.
  9. Transfer sol’n into 125 ml flask
  10. Clean filter unit by rinsing beads out and then vacuum washing 3X with distilled water.
  11. Allow to air dry (filter can be used until tears are visible).


  1. Use rinsed 250 ml flask from #1 above as syringe stand.
  2. Add 20??l of TEMED and 600??l of 10% APS to gel mix, swirl well to mix.
  3. Pour mix into 60cc syringe with ground off needle.
  4. Angle plates with use of rack and hands and allow mix to gravity feed between plates.
  5. Stop and remove any bubbles.
  6. Place combs so bottom of oval lines up with top edge of smaller glass.
  7. Pour sol’n to cover combs.
  8. Remove clamps and slowly pull out bottom spacer.
  9. Reposition top clip on each side inward slightly.
  10. Tilt leftover sol’n in flask to indicate when plates have polymerizied.
  11. Clean syringe, etc. so gel doesn’t polymerize.

Gel Setup

  1. Prepare buffer while gel is polymerizing. In a grad. cylinder, dilute 25 ml of 20X Glycerol Tolerant Buffer (50 ml 10X TBE) for a FV of 500 ml in ddH2O for upper buffer chamber.
  2. Also, in a grad cylinder dilute 10 ml of 20X Glycerol Tolerant Buffer (20 ml 10X TBE) for a FV of 200 ml in ddH2O for lower buffer chamber.
  3. After polymerization, remove clips, rinse plates, remove combs and rinse excess acry away.
  4. Close upper buffer chamber drain on right side of apparatus.
  5. Place gel in “box”, clamp into place, put buffer in upper chamber.
  6. Put buffer in lower chamber making sure there are NO bubbles under gel.
  7. Puff bubbles and urea from between plates.
  8. Place combs so tips are at surface but not into gel.
  9. Run at 65 watts for 5-15 minutes.

Sample Preparation and Loading

  1. Radioactive components will be underlined from this point on.
  2. Divide samples and place approx. half in 70??C heat block for 2-10 minutes.
  3. Mark gel for samples (4 lanes each) and comb meeting.
  4. Load 3??l of samples L to R, alphabetically (A,C,G,T). Put tips in solid radioactive waste container!
  5. Run at 65 watts while other samples are in the heat block. Do this so a) makes gel asymmetrical b) gets samples into gel before diffuse.
  6. Load other samples. Put tips in solid radioactive waste container!
  7. Run total of 2′ 45″.
  8. Prepare 80 g NaOAc into 200 ml (not FV) ddH2O. Stir till dissolved on stir plate H2O bath with some heat.
  9. One hour before gel is done, add NaOAc sol’n to lower buffer chamber. Helps slow down front and compresses bands.
  10. Cut a piece of Whatman paper 43 x 34 cm & roll up into pipet cylinder lid.

Take Down and Drying

  1. Prep: have two green test tube racks set aside for top plate (treated), rolled paper, water bottle, and spatula.
  2. Drain upper chamber into holding well.
  3. Disconnect plates.
  4. Let plates drain into lower chamber — bottom edge is hot!
  5. Rinse off edge in sink.
  6. On bench, remove combs.
  7. Separate plates with spatula placing top plate gel side up on racks.
  8. Squirt water around sides and top of gel on plate.
  9. Position paper over the gel and place on gel from center.
  10. Smooth firmly from center with hands.
  11. Water assists in gel rolling up onto paper.
  12. Transfer gel/paper to dryer.
  13. Layer in the following manner (bottom to top): Mesh, filter paper to protect mesh, gel/paper (gel side up), saran wrap (must completely cover gel/paper), plastic cover, gasket.
  14. Dry for 2 hours at 80??C.
  15. Let cool 15-30 min before trying to remove from dryer.

Film Exposure and Development

  1. Remove gel/paper from dryer.
  2. Take gel, cassette and Biomax MR film to dark room.
  3. Lock door and turn on outside “in use” light.
  4. Place film in cassette with notch in upper right corner.
  5. Place gel face down with top of gel at top of film, close the cassette.
  6. Steps 3 and 4 allow gel to be read with the notch in the upper left corner.
  7. Leave at room temperature in drawer for 20 hr for S35, 20 hr for P33, and 8 hr if P33??is fresh.
  8. To develop, place one edge against side of film developer.
  9. Make sure write down what size film was developed on sheet.


  1. Rinse clips, spacers, and combs. Allow to air dry.
  2. Rinse plates well but not necessary to soap wash. Allow to air dry.
  3. Put everything back in proper place when dry.

Reaction Tube Color Code:??

A = black (all colors)
C = red (crimson)
G = green
T = blue (teal)

Seqencing Reactions Using Amersham Thermosequenase Kit

  1. a) Get full tub and bucket of ice, b) get isotope out if in -20??C
  2. Get dGTP or dITP master mix buffer and reaction buffer out and thaw on ice
  3. Thaw primers on ice, they need to be 2 pmol/l
  4. Label 4 PCR tubes per sample in the following order and color code:??
    Base Letter Marker Color Description
    A black all colors
    C red crimson
    G green ??
    T blue teal

  5. Label 4 colored tubes for termination mixes and??_X_??colored tubes for sample reaction mixes
  6. Get dNTPs out to thaw
  7. Do termination mix calc. using kit quick card as preparing for (n + 1) samples
  8. Do reaction mix calc. (normally 1??l of mini prep is used, so adjust H2O)
  9. Transfer 2.5??l termination mix to each appropriate tube (left to right)
  10. Transfer 4.5??l reaction mix to each appropriate tube (front to back) and mix sol’n with pipet tip

    Tub diagram:

    T T T T T
    Hot G G G G G
    bases C C C C C
    A A A A A
    Samples: 1 2 3 4 5

  11. If using “Bonnie” don’t need to overlay with oil
  12. Thaw stop sol’n at end of cycles and add 4??l to each tube using single tip
  13. Spin pulse all samples
  14. Store at -20??C in refrig freezer


  • Make sure all “hot” tips and bullets get put into the radioactive solid waste
  • In our experience, the reaction will not work if the buffers are switched
  • It is very easy to get confused, allow yourself plenty of time to set up and mix the reactions; they can be done the day before and froze

Library Screen

Day 1

  • Start overnight of LE392 in LB with 10mM MgSO4

Day 2

  • Make 12 plates for lifts using 10,000 plaques per plate dilution
  • Combine 10??l of dilution to be plated with 100??l of LE392 in 1.7 ml bullet
  • Warm (no shaking) at 37??C for 15 min
  • Add phage/LE392 to 3 ml lambda top agarose cooled to 50??C
  • Working quickly, vortex for 10 seconds and pour all 3 ml onto a pre-warmed (37??C) NZY or LBM plate; rock plate to spread agarose out over entire surface
  • Once plates are solidified, incubate plates at 37??C for 6 hours
  • Prepare probe following probe labeling protocol
  • After 6 hours, place plates in single layer for 1 hour at 4??C to chill
  • While chilling, label nylon circles with pencil or sharpie
  • Prepare denat. and neutral. filter paper/saran wrap stations
  • Have small amount of 2xSSPE in square glass dish for rinsing
  • Have filter paper for drying circles and filter paper size of crosslinker to put circles on (re-use both of these)
  • After 1 hour, do lifts with writing side up marking membranes with 18G needle (can re-use also) (!!!! SAVE PLATES!!!!)
  • Denature for 5 min from start of the first circle (writing side up)
  • Neutralize for 5 min from start of the first circle (writing side up)
  • Rinse in 2x SSPE and lay plaque side up on extra filter paper (writing side down)
  • Let dry until no pooled liquid is visible on surface (do not overdry)
  • Crosslink on auto setting in crosslinker
  • Transfer circles to seal a meal bag with circles back to back (plaque side out)
  • Add 15 ml hybridization solution warmed from 37??C to 42??C
  • Add 200??l (133??g/ml) of 10 mg/ml sonicated salmon sperm DNA which has been heated for 5 min at 95??C
  • Don’t seal, just lay in hybrid oven at 42??C for at least 5 min
  • Add amount of probe determined, mix, and seal bag
  • Transfer bag(s) to zip lock bag
  • Put in hybrid. oven and rock o/n at 42??C noting the time started

Day 3

  • Note time hybridization stopped
  • Cut bag and pour solution into liquid wastes using kimwipe to wipe edge
  • Remove circles to small amount of 0.2x SSPE/0.1% SDS (wash sol’n) in small glass dish
  • Hand agitate and pour sol’n into liquid waste, wipe edge with kimwipe
  • Add 300-400 ml wash sol’n
  • Agitate 20 min at 50??C in water filled hot shaker
  • Pour sol’n into liquid waste, wipe edge with kimwipe
  • Wash two more times (3 x 20 min washes total)
  • Pour #2 and #3 wash sol’n into sink rinsing corner with hot water
  • Transfer circles to filter paper
  • Stretch and tape saran wrap over film cassette template
  • Lay filters on wrap with writing up
  • Stretch and seal wrap, trim edges
  • Put in cassette with plaques down and intensifying screen below
  • Put orientation labels on saran wrap to align circles with film
  • Put film(notch in upper right corner if notched) between screen and circles
  • Expose 4-24 hour at -70??C depending on counter detected signal

After Film is Developed

  • Align labels between film and circles
  • Mark needle holes with fat sharpie
  • Trace over circle label, circle possible positives, and label film with date, time exposed and probe used
  • For positives, align appropriate plate on film using needle marks
  • Check to see if positive on film lines up with plaque on plate
  • If so, pull plug of plaque and put in 1 ml of SM in 1.7 ml bullet with 1 drop of chloroform, store at 4??C
  • Rescreen

Library Titer For Zap Cdna

  • Want the dilution that gives 10,000 plaques per 100 mm diameter plate
  • Keep all dilutions made from the library for later use

Day 1

  • Streak LE392 onto a NZY or LBM plate and grow o/n at 37??C

Day 2

  • Start an o/n from the fresh plate of LE392 in LB with 10mM MgSO4
  • Make the following dilution series:??

  • Combine 10??l of dilution to be plated with 100??l of LE392 in 1.7 ml bullet
  • Warm (no shaking) at 37??C for 15 min
  • Add phage/LE392 to 3 ml lambda top agarose cooled to 50??C
  • Working quickly, vortex for 10 seconds and pour all 3 ml onto a pre-warmed (37??C) NZY or LBM plate; rock plate to spread agarose out over entire surface
  • Once plates are solidified, place at 37??C o/n

Day 3

  • Count plate(s) that show 30-300 plaques per plate
  • Back calculate to determine library concentration
  • Determine dilution to use for 10,000 plaques per plate

Probe Counter Beckman Across From Denells Lab

  • Remove cover from printer
  • From center tower the machine goes clockwise
  • From left to right put in stop, sample(s), and user tag on #8 (inserts into holder only one way)
  • Close door and push auto (green)
  • Will print report automatically
  • When finished put printer cover back on so it does NOT cover fan vents

Probe Labeling Using Decamer Oligos

Standard Reaction

Final Volume of rxn is 50??l. Make sure water added to DNA in step 1 is adjusted for the amount of isotope used in step 2. Components in -20??C box: “oligo labelling”.??

Step 1: 1??l DNA (approx. 100ng/l)
?? 29??l ddH2O
?? ?? – heat 5 min at 95??C
– cool on ice
– spin
Step 2: add??10??l 5X decamer buffer
?? 2??l BSA (10mg/ml)
?? 2??l dNTPs minus dCTP (0.5 mM)
?? 0.5??l exo(minus) Klenow
?? 5??l P32??dCTP (fresh)
?? 50??l total volume

  • Transfer tube to 37??C heat block for 30-45 minutes
  • Add 50??l TE(10/1) to reaction and transfer to a dried (2 min at medium in clinical) G-50 column
  • Spin 2 min on medium
  • Add 100??l TE to column to rinse
  • Spin 2 min on medium
  • Transfer sol’n to new labelled tube and estimate volume
  • Transfer 1??l to small piece of filter paper for counting and allow to dry
  • Count probe on Johnson’s machine with user #6 card(see below)
  • Store probe in refrigerator -20??C in plexiglass
  • Before use add 1/5 volume of 1M NaOH to denature probe (do not boil)
  • Let sit 5 min
  • Add 2×107??counts per roller or bag — make sure amount has been adjusted for NaOH volume addition)
  • Probe needs to be at least 1×105??to use
On probe counter printout, calculate the following:
counts/l total counts l for 2x107counts/bag 1M NaOH to add adjusted vol. for 2x107counts/bag

Expand Rocheboehringermannheim Long Template Pcr

Master Mix I Master Mix II
template enzyme
primers buffer
dNTPs ddH2O to 25??l
ddH2O to 25??l


  • Preheat PCR machine to 94??C using instant key
  • Mix the above two for 1 reaction F.V. 50??l
  • Use thin wall 0.2 ml PCR tubes
  • Vortex to mix thoroughly
  • Spin pulse
  • Overlay with 30??l oil even with hot bonnet
  • Master mix I can be mixed in thin wall tube when few samples
  • Master mixes of I or II can be done for large # of reactions