Adapted from Bohlander et al. Genomics 13 (1992).
(modified by D. Schubeler, L. Loo)

Genomic DNA will be randomly primed with a sequence tagged oligonucleotide for 2 cycles.  This will create random genomic products with a specific tag at both ends. These products will then be amplified and labeled with a sequence specific dye primer. The protocol has been optimized for DNA amounts ranging from 1 to 50 ng.  Matt's notes: Template DNA template which exceeds the capacity of Sequenase (in Part A) would carry over and unnecessarily contaminate the PCR reaction in part B.  However part B should be able to handle much larger quantities of sequence tagged substrate than is produced in the Part A reaction.

For two color arrays an equal amount of control DNA should also be amplified in parallel to the experimental DNA.

Material

Protocol

Part A. DNA fragmentation and Priming reactions

1. Quantify genomic DNA by spectrophotemetry of fluorometry and dilute in T.E. to 40 ng/mL.
2. Create a restriction digest master mix by multiplying the following recipe x the number of unknown plus control DNA samples (+10% for pipetting error:
Reaction 1:

0.7 µL 10x Sau3a buffer
0.07 µL 10 mg/mL BSA
0.35 µL Sau3a enzyme
5 µL H2O

3. Setup the digest by adding 6 µL of this mix per DNA sample (+ controls) in PCR tubes.

Add 1 µL DNA (40 ng) to each tube.

Digest DNA then heat inactivate enzyme on PCR machine:
(37ºC x60', 65ºC x10') x1
(4ºC hold)

4. While the genomic DNA is digesting set up master mixes for Reactions I, II, and III by multiplying the following recipe by the number of samples (and controls) + 10% for pipetting error:

5. Add 3 µL of Reaction 2 to each tube. Start the Part A PCR program. Note: Reaction 3 and 4 mixes will be added as the machine is cycling as explained below:

Part A -- PCR machine parameters:
(94°C 2’, 10°C 5’ {add reaction 3 or 4 here}, 8’ ramp to 37°C, 37˚C 8’) x2
(4ºC hold) x1

6. Let the machine finish heating to 94°C x 2 min. When it then cools to 10˚C add 5 µL of Reaction 3 mix to the sample. The machine should then slowly ramp to 37°C over 8 min. and then hold at 37°C for an additional 8 min to complete the first cycle.

7. During the second cycle (again at 10ºC) spike each sample with 1.2 µL of Reaction 4 mix.
8. When the cycling is complete add 43.8 µL dH2O to each sample to bring their final volumes to 60 µl.

Part B. PCR amplification with Cy-Dye primers:

The part A reactions produced enough DNA product for 2 part B reactions. For example 2 arrays could be set up with a dye swap. The FHCRC human BAC array has internal duplicates so performing duplicate arrays is redundant. If a single reaction will be performed Cy3 should be reserved for the reference (control) DNA since this fluorochrome has more autoflourescense and a more restricted linear range. (i.e. curvature at low intensities in log ratio plots.)

1. Assuming only 1 array will be run per sample then create master mixes for Cy5 primers by multiplying the following recipe by the numbers of samples (+ 10% for pipetting errors). An equal amount of Cy3 mix should be made for the reference DNA.

Round B PCR
30 µL Round A DNA product
20 µL 10X Thermo Pol PCR Buffer
2 µL 25 mM dNTP
4 µL 100 µM Primer B Cy5 (sample) or Cy3 (reference DNA)
2 µL Ampli Taq
0.1 µL Pfu Turbo
142 µL dH2O
Total 200 µL

Matt’s note: With this recipe primers and dNTPs are stoichiometrically balanced if the mean product is 250 bp. The total amount of dNTPs is 15 µg/200 µL.

2. Divide the each primer mix into 4 PCR tubes (50 µL each). Start the Round B PCR program and place the samples on the machine when it reaches 94˚C.

Part B -- PCR machine parameters:
(94˚C 3’) x1
(94˚C 30”, 40˚C 30”, 50˚C 30”, 72˚C 1’) x35 cycles
(4ºC hold) x1

3. To check if the PCR reaction went well run 3 µL on 1% agarose gel. A smear of DNA should be present between 0.5 – 1 kb. (It can be saved to run with step B.4.d).

Part C. Preparation of Hybridization Mixture:

Caution: The product of Part B is a potent source of contamination for future amplifications and should be handled with the same care as any PCR product (gloves, filter tips, benchkote paper). Consider resticting the work area and equipment while handling this material.

1a. To remove unincorporated nucleotides use a Sephadex G-50 spin column (1a). Alternatively, a PCR cleanup column (1b). Sephadex columns can be performed according to Molecular Cloning (Vol.3 A8.29), or with a Roche DNA QuickSpin column. First equilibrate the column with T.E. Next pool 4 identical PCR reactions and run them through a single column into an amber 1.5 mL Eppendorf tube.

1b. As an alternative to sephadex columns clean up the PCR product with QiaQuick PCR cleanup columns. Follow the manufacturer's directions. In summary: Open the PCR tubes and add 250 µL of solution PB to each. With a P200 pipettor mix the DNA and PB from 2 PCR tubes and combine them in a single QiaQuick column. (Theoretcally the 4 PCR tubes per sample yield 15 µg of PCR product - based on the amount of dNTPs, and each QiaQuick column has a capacity of 10 µg). Repeat this with an identical PCR sample (labeled with the same dye) to pool the products of two PCR tubes in each column. Pool two identical PCR reactions (labeled with a single dye) to make 100 µl DNA. add 500 µl of PB. Centrifuge for 30" - 60". Wash with 750 µL PE and spin. Elute DNA by placing column into an amber 1.5 mL Eppendorf tube, add 50 µL EB and spin. (Optional - add 25 uL EB to column and repeat spin.)

2. Pool identical sample (labled with same dyes). Save a 3 µL aliquot for analysis on a 1% agarose gel.

3. Add to each pooled PCR product add:

50 µL Human Cot-1 DNA (1 µg/µL)
10 µL tRNA (10 µg/µL)

4. Concentrate samples on a Microcon YM-10 concentrator (MW cutoff ~ 10 kDal, 17 bp). (Alternatively YM-30 concentrators can be used which have the advantage of being quicker and they eliminate primer dimers - MW cutoff ~ 30kDal, 50 bp). Spin at 14,000g until volume is ~25 µL. Spinning to smaller volumes will help to eliminate contaminating small molecules and will speed up the next step.

5. Pool each experimental sample (Cy3 labeled) with an equivalent amount of control sample (Cy5 labeled) and dry down in speed vac to a volume of <5 µL (not too dry!).

(Ref: Molecular Cloning A8.12)

The major considerations in using alcohol to precipitate DNA are:

1. Temperature: -20°C is optimal, but 0°C can be used for >20 ng/mL
2. Amount: For small amount of DNA (<100 ng, i.e. too little to reliably see a pellet) the use of glycogen (1 µL of a 20 mg/mL stock, (Roche 10901393001) will increase yield and allow visualization of the pellet.
3. Time: Optimal precipitation requires >1 hr at -20°C.
4. Speed of centrifuge: High speeds are important for small amounts or small oligos. Typically they should be spun at >12,000 rpm (> 12,000 g) in a microcentrifuge.
5. Mg++ [10 uM] helps pellet oligonucleotides (<100 bp). Otherwise Mg should be avoided because it may activate DNAses.
6. Avoid co-precipitates: EDTA (>10 mM) and Ca (>1 mM) will precipitate at the concentrations indicated in alcohol solutions. SDS will also precipitate when a salt other than NaCl is employed.
7. Choice of alcohol: Isopropanol has the advantage of requiring less volume. Typically 0.6 vol to 1 vol. of isopropanol is added to 1 volume of DNA/salt soulution. The higher amount is useful for smaller RNA fragments. In contrast, 2 volumes of ethanol is typical.) However, isopropanol has the disadvantage of coprecipitating more salts and is less volatile (so it is slow to air dry) compared to ethanol.
8. Choice of salt:

NaOAc, pH5.2, [0.3M]: This is the standard. (10x Stock concentration = 3 M)

NaCl [0.2M]: Useful if SDS is included because it is more soluable in NaCl (Stock conentrations vary 1M - 5M)

NH4OAc [2 - 2.5M]: Less coprecipitation of dNTPs, good for purifying oligos. However, NH4+ ions inhibit polynucleotide kinase).

LiCl [0.8M]: Soluble in a higher concentration of ethanol which is useful for the precipitation of RNA. However Cl- inhibits the initiation of protein synthesis and RNA will display varying solubility based on size.

Solutions

PBS
Cell lysis buffer: 10 mM Tris pH8, 100 mM NaCl, 25 mM EDTA, 0.5% (w/v) SDS, 0.5 mg/mL proteinase K
Fixative: 1 volume glacial acetic acid + 3 volumes methanol
Trizol reagent
75% EtOH made with RNAse free H2O
100% EtOH
Phenol: Buffered to pH=8 with Tris.
Chloroform: Stabilized with isoamyl alcohol.
T.E.: 10 mM Tris pH8, 1 mM EDTA.

Note
This protocol prepares genomic DNA from cryopreserved leukemia cells. It assumes a starting volume of ~1.8 mL and uses only 1/2 of this volume. The remainder is frozen again. The DNA preparation protocol does not contain a RNAse step. It may be necessary to add this if a significant amount of RNA contaminates the DNA. It also saves an aliquot of RNA from each sample, partially purified by the Trizol protocol. A third aliquot is used to fix cells for cytology.

Procedure

1. Place freezer boxes on dry ice.
2. Place vial in a 37ºC water bath just long enough to thaw.
3. Mix gently and transfer 0.9 mL of cells to a 1.5 mL tube. The remainder of cells should be placed in a control rate freezing jar and placed at -80ºC. Transfer back to the original freezer boxes at -80ºC. Make a note in the MPD Frozen Samples that 1/2 of the sample was used.
4. Spin at 600g. x 2 min.
5. Pipet off supernatant and discard. Do not use an aspirator because lysed cells may release viscous DNA which is adherent to the cell pellet.
6. Dislodge cell pellet by flicking tube and resuspend in 0.2 mL cold PBS.
   • DNA: Transfer 0.1 mL to a tube with 0.4 mL of cell lysis buffer and continue with step 7, below.
   • Fixed cells: Transfer 20 µL to a tube with 0.5 mL fixative (1:3, HOAc:MeOH). Spot 1 drop onto a glass slide and stain with Hema3 stain. Transfer remainder to a 1.8 mL cryo vial and store at -20ºC. List fixed cells and histo slide in MPD freezer database and Human Tissue Reports.
   • RNA: Spin remainder of cells. Discard supernatant. Add 0.6 mL Trizol. See Trizol protocol for details on RNA preparation. Store RNA in 70% EtOH at -20ºC. List RNA samples in MPD freezer database and Human Tissue Reports.
7. Place DNA in lysis buffer with proteinase K at 50ºC x 1hr, and mix periodically.
8. Add 2 vol. (1 mL) of phenol. Mix gently at 4ºC for 1 hr. to overnight. Transfer aqueous phase to a fresh tube.
9. Add 1 vol. (0.5 mL) of CHCl3, mix gently to remove phenol. Transfer aqueous phase to a fresh tube.
10. Ethanol precipiptate by adding 2 vol. (1 mL) EtOH, mix and incubate -20ºC for at least 1 hr. Spin at 12,000 rpm to pellet DNA and discard supernatant.
11. Wash with 70% EtOH x2. Air dry. Resuspend in 0.1 mL T.E. Store at -80ºC. List DNA samples in MPD freezer database and Human Tissue Reports.

Materials

Electroelution gel box and elution block
0.25x TBE buffer
5 M KOAc with BPB (bromphenol blue)
100% EtOH
70% EtOH
T.E. buffer (10 mM Tris, 1 mM EDTA, pH 8)

Procedure

1. Run your DNA fragment on a preparative agarose gel
2. Add 350 mL of 0.25x TBE buffer into electroelution box. Buffer should cover the ports on the elution block but should not cover the top of the block. Remove any bubbles from the elution block by flushing the tubes with TBE buffer. Do not use higher concentrations of TBE buffer.
3. Using a pipetman with a gel loading tip add 40 µL of 5M KOAc/BPB to the sloping tunnel of the device, starting at the bottom of the well, then gradually withdrawing the pipet tip as the well fills. The vertical tunnel should remain filled with TBE buffer.
4. Cut gel fragment containing DNA band into 2 mm wide strips.
5. Add 2-4 mm gel strips to the slot in the elution device, closest to the elution port.
6. Electroelute at 100v. x50 min. for a 1 kb fragment.
7. Using a gel loading tip remove 50 uL of the blue solution from the device.
8. Let the solution settle and remove the remaining solution to bring the total volume to 100 µL.
9. Precipitate the DNA by adding 200 µL of 100% EtOH. Incubate at -20ºC x30 min.
10. Spin at 13,000 rpm, 4ºC.
11. Wash 3x with ethanol:
    1. Add 1 mL 70% EtOH, mix gently.
    2. Spin x2 min. at max. speed. Decant off supernatant.
12. Air dry.
13. Resuspend in a minimum volume of T.E.
14. Check concentration and quality of product by running 1 µL of DNA on a test gel with a DNA size standard.

M. Fero 11/05  •  Adapted from Turner's protocol [Promega bought Turner Biosystems in 2009. Promega references some Turner Biosystems items on their website]

Materials

DNA: Höchst 33258 (High range solution):  (Invitrogen H1398 or Sigma B1155) 1µg/mL in 1x TNE

• Filter and store at 4ºC in an amber bottle.
• 10x TNE stock:  100 mM Tris, 2 M NaCl, 10 mM EDTA, pH7.4.  Store at room temp.

RNA:  Ribogreen 200x (Invitrogen R-11491) detection reagent.  Store dessicated at -20ºC.
Turner Designs TD-360 Fluorometer

• DNA: Long UV LED (underneath) and filter set (inside chamber)
• RNA: Blue LED and filter set

RNA or DNA standards (100 ng/µL)
TE pH 8 (DNA), pH 7.5 (RNA)

Discussion

Fluorometry utilizes fluorescent dyes which specifically bind DNA or RNA.  It requires a negative control (to set the zero point on the fluorometer) and a standard of known concentration.  The fluorometer shines light on the sample (excitation) and then measues level of fluorescent light being emitted to the side (at a 90º angle) of the excitation light beam.  The fluorescent dyes are relatively specific to nucleic acids as opposed to protein and other cellular components.  The fluorescence of the dyes increases when they bind nucleic acids.  Fluorometry is about 1,000x more sensitive than spectrophotometric absorbance (i.e. measurement of A260) and less susceptible to protein and RNA contamination.  However it also does not give a crude measurement of purity (like an A260/A280 ratio) nor does it assure that the DNA or RNA is not degraded (e.g. like size determination by gel electrophoesis).  Do not use glass (spectrophotmetry) cuvettes in a fluorometer because the frosted glass on the side of the cuvette interferes with detection of fluorescent light.

Detection Ranges

Procedure

(High range assay solution)

1. Insert the correct LED and filter set in the fluorometer for either RNA or DNA.
2. Turn on the Fluorometer to warm up for 5 min.
3. DNA:  Remove Höchst (high range) solution from the refrigerator.  Dilute Höchst 1/10 (v/v) in 1x TNE if a low range assay is being performed.
   • Large cuvettes require 1.5 mL/sample
   • Mini-cuvettes need 100 µL/sample.Aliquot 1.5 mL (large cuvette) or 0.1 mL (microcuvette) of either Höchst or Ribogreen reagent into 1.5 mL eppendorf tubes.
     
4. Add 1.5 µL of DNA or RNA to each tube and vortex.  In addition to samples this must also include a known standard and a TE or H2O control.
5. Using narrow (gel loading) tips pipet the TE or H2O control in a cuvette, close the chamber's lid and press the "Blank" button.  Press "1" when prompted to save the data.  Open the chamber's lid and transfer the control mix back to its original Eppendorf tube.
   
6. Transfer the DNA or RNA standard to the cuvette and press the "Calibrate" button.  When prompted enter the (undiluted) concentration of the standard (i.e. 100) plus the "Enter" button. Press "1" when prompted to save the data.
7. Repeat the zero and calibration steps (6 and 7).  This is important to avoid negative values at low concentrations.
   
8. Measure each RNA sample and record the concentrations.
   

Note:  The procedure for "Low range" assays is identical but it uses the detection reagent diluted 1/10x the cocentrations listed above to minimize background autofluorescence.

Measuring DNA concentration with the Nanodrop Spectrophotometer

v.1 10-31-2011
A. AlKhinji

Materials

• DNA in TE buffer (10 mM Tris pH8, 1 mM EDTA)
• TE buffer for blanking machine
• P20 micropipettor + tips
• H2O squeeze bottle and Kipwipes (at machine)

Procedure

1. Sign log book
2. Login to computer: (Adjacent lab P.I./room)
3. Launch software: ND-100 v.3.7.1
4. Wash the lower (sensor) pedestal and the upper (lid) pedestal with H2O spray bottle, then wipe both pedestals with a Kimwipe.
5. Add 1-1.5 µL H2O. → Select "Initialize". Wipe pedestals clean.
6. Add 1-1.5 λ TE → SELECT "Blank". Wipe pedestals.
7. Repeat the following for each sample:
8. Enter Sample #
9. Add 1-1.5 µL DNA sample → Select "Measure".
10. Wipe pedestals.
11. Select "Show report" > Save report and export as Tab-delimited text file.
12. When done: Wash the upper and lower pedestals with with H2O and wipe to dry.
13. Select "Exit".
14. Log off computer.

Comments

The Nanodrop will display an absorbance spectrum for each sample as it is being measured. DNA should have an absorbance peak centered at a wavelength of 260 nm (A260). The ratio A260/A280 should be ~1.95. The presence of organic solvents (e.g. phenol) may lead to a spuriously high A260/A280 ratio (> 2). Proteins will have a peak absorbance at 280 nm, so protein contamination will lower the A260/A280 ratio. Protein does not absorb as strongly as DNA so even a modest reduction in the A260/A280 ratio (e.g. 1.8) may represent a high level of contamination. The extinction coefficient is a factor that converts the peak absorbance to concentration. For DNA the extinction coefficient is 50 (ng/µL DNA) / A260. The nanodrop has a broad linear range. Accuracy drops off (error > 10%) for concentrations < 4 ng/µL and > 4000 ng/µL.

Notes
Stocks should be handled with care to ensure that they are not contaminated with PCR product or plasmid DNA that might be prevalent in the laboratory. For this reason they are handled only in the pre-PCR area with gloves, aerosol resistant tips, and DNAse/contamination-free solutions.

Oligo concentrations are typically expressed in terms of moles, not weight (µg). The extinction coefficient, for converting absorbance (A260) to concentration (µM), varies according to the base composition of the oligo. In this protocol, the MPD is used to record the oligo sequence, to calculate its extinction coefficient and concentration, and to note the location of the freezer stock solution.

Materials

• DNAse free H2O or Tris 10mM, pH=8
• 1.5 mL Eppendorf tubes
• Aerosol resistant pipet tips
• Spectrophotometer
• Oligonucleotide stock (from manufacturer)

Procedure

1. Turn on spectrophotometer.
2. Take bag containing oligo to pre-PCR room. Dump tube on bench, without touching tube or inner surface of bag.
3. Don gloves. Place tube in pre-PCR rack.
4. Add 10 µL of H2O or 10mM tris per nMole of stock primer (quantity according to the oligo maker). Vortex.
5. Dilute Oligo 1:50 v:v by adding 10 uL to 490 uL H2O. Also aliquot 500 uL H2O as a blank.
6. Using a glass cuvette (not plastic) blank the spec. using the H2O. Measure A260 of the oligo.
7. Find or enter the oligo name in the MPD on the Oligo Details layout. Enter the A260 value and dilution factor (50).
   
   

   A260
   Dilution Factor	50
   Concentration (µM)	0.00

   
   Record the concentration of the oligo in your notebook.

8. In the pre-PCR room don fresh gloves and write the concentration of the oligo on the stock tube.
9. In a 1.5 mL tube, dilute the primer stock down to the working concentration of 10 uM with H2O or 10 mM Tris, pH8.
10. Place the remaining oligo stock at -80ºC, and record it's location in the MPD.

SYBRGreen Q-PCR in the ABI 7700

Materials

• ROX Passive Reference Dye 25 µM, 50x (Invitrogen, Cat. No. 12223-012)
• SYBR Green I, "10,000x" concentrate (Molecular Probes, Cat. No. S7563)
• SureStart Taq polymerase 5 U/µL (Stratagene, Cat. No. 600280)

Solutions

• SYBR Green 1:100 dilution (50 µL SYBR Green concentrate, 5 mL DMSO, store aliquots at -20ºC in dark tubes in a dessicator)
• SYBR Green 1:2000 solution (0.5 mL of SYBR Green 1:100 dil., 1 mL glycerol, 10 uL Tween, 8.5 mL H2O, store at -20ºC in dark tubes)

PCR Reaction Setup
1. Create a master mix by multiplying this recipe by the number of reactions (add 10% for pipeting error):
2 µL Primer-1
2 µL Primer-2
2 µL KG-1
2 µL KG-2
5 µL 1:2000 SYBR Green solution
0.4 µL ROX Dye 25 µM
0.1 µL SureStart Taq
6.5 µL H2O
2. Setup individual reactions in PCR tubes or plates with optical caps or optical tape:
20 µL Master Mix
1 µL DNA (~ 50 ng genomic DNA)

Machine Setup
1. If the machine status reads "Idle" but a user still has not remove their samples then "Save" the current file to preserve the users data before shutting down.
2. Shut down both the machine and the computer. Turn on the detector first then boot the computer.
3. Launch SDS v1.9 software (don't use v1.7).
4. Create a new template:
Dye Layer = SybrGreen
Sample Type = Sample type setup. Choose SYBRG for UNKN, NTC. Deselect quencher.
Select wells with samples. Set sample type to UNKN (or NTC).
5. Program thermocycle conditions
95ºC 5' Hold (Stage I)
95ºC 15", 60ºC 30", 72ºC 30" (x40) (Stage II - this stage is emperic)
60ºC 15" Hold (Stage III)
95ºC 15" Hold (Stage IV)
Select Stage IV and set ramp time to maximum (19:59).
Select "Show Data Collection..."
Select and delete the documents for stage III and IV.
Select to add documentation of the Stage IV ramp.
6. Select "Show Analsis..." > Instrument > Diagnostics > Advanced Options
Select: Set 7700 exposure time for plates 25 (caps), or 10 (film).
Reference = Rox (unless no reference is used)
7. Save the template to the "Fero Lab" folder.
8. Place plate in sample block, close lid and shut door. Select "RUN".

Saving and processing data
9. When the machine has finished cycling the status will read, "Idle". Save the file again before quitting.
10. To save dissociation curves: select File > Export... > Multicomponent...

Materials

Procedure

1. Mix Prehybridization Solution until dissolved. Incubate filters at 42°C in Prehyb solution in either the glass bell jar or a Seal A Meal bag for 1-4 hrs.
2. Turn on heater block to 100°C.
3. Transfer probe to 1.5 mL eppendorf tube with a pinhole in the cap and add H2O to bring the total volume to 150 µL. Denature the DNA by heating to 100°C for 5 min.
4. Transfer probe to the filter and chamber. Mix well with the Prehyb solution. Reseal the bag or put a lid and weight on the bell jar. Incubate with shaking at 42°C overnight.

Washing Filters

Materials

• 20x SSC (3M NaCl, 0.3M Citrate, pH 7.4)
• 65°C shaking water bath
• Whatman 3M paper
• Saran wrap, florescent stickers
• Autoradiograph casette and intensifying screens
• X-ray film (Kodak BioMax 8 x 10", Cat 829-4985)

1. Discard the radioactive Hybridization solution into absorbed liquid waste container. Rinse the filter with 1x SSC, 0.1%SDS and discard.
2. Wash the filter in 1x SSC, 0.1% SDS in a shaking water bath for 1 hour. Meanwhile allow the water bath to heat to 65°C. Change wash solution 2x during the hour.
3. Wash the filter in 0.1x SSC, 0.1% SDS at 65°C for 10 to 20 minutes. Meanwhile monitor the CPM on the filter with a Geiger counter. Stop washing the blot when the counts are reduced to 2 - 4x background.
4. Blot the filters on Whatman 3MM paper. Wrap the filter in Saran wrap and place glow in the dark stickers for orientation. In a dark room place the filter in an autoradiograph casette then a piece of X-ray film, and then an intensifying screen. If a weak signal is anticipated do this in complete darkness (with the safe lights off).
5. Place the film casette at -70°C. For cloned DNA and high S.A. probes expose for 2 - 4 hours. For mammalian genomic DNA Southern blots expose the film for 18 - 72 hrs.
6. Remove the film casette and allow it to warm for 30 min. Remove the film in the dark room (for weak signals do this with the safe lights off) and develop in a processor or in developing tanks as follows: X-ray developer: 2 min. Water wash 10 sec. Fixer: 2 min. Water wash 5 min. Air dry the film.

Final Concentration

• 5x SSC
• 25 mM NaPO4, pH6.5
• 5x Denhardt's Solution
• 0.25 mg/mL Torula RNA
• 50% (v/v) Formamide

Stock Solutions

• 20x SSC: 3 M NaCl, 0.3 M NaCitrate, pH7.0
• 50x Denhardt's: 1% (w/v) BSA, 1% (w/v) PVP-40, 1% (w/v) Ficoll-400
• 10 mg/mL Torula RNA (boil to dissolve)
• 1 M NaPO4, pH 6.5

Preparation

To make 1 L. of Stark's Buffer mix:

• 250 mL 20x SSC
• 25 mL 1M NaPO4, pH6.5
• 100 mL 50x Denhardt's solution
• 25 mL 10 mg/mL Torula RNA (boil just prior to adding)
• 122.5 mL H2O
• 500 mL Formamide

Reagents

• Lysis Buffer: (10 mM Tris pH8, 100 mM NaCl, 25 mM EDTA, 0.5%SDS), store at room temp.
  10 mg/mL Proteinase K: 100 mg (Roche) + 10 mL buffer (10 mM Tris pH8.0, 20 mM CaCl2, 50% (v/v) glycerol), store at -20ºC.
  Lysis buffer + 1 mg/mL proteinase K: 50 mL Lysis buffer (above) + 0.5 mL 10 mg/mL proteinase K, store at -20ºC.
  TE: 10 mM Tris pH8, 1mM EDTA
• Phenol: Molecular biology grade. Equillibrate 1x with Tris pH8, and then 1x with T.E. Store at 4°C.
• Chloroform: Fresh choloroform + 1/20 vol. isoamyl alcohol.
• Absolute ethanol.

Procedure

1. Cut 3 mm of toe or tail or toe tips from 7-10 day old mice into 1.5 mL microcentrifuge tubes.
2. Digest tail biopsy by adding 0.7 mL lysis buffer + proteinase K and incubate for 4-16 hrs at 37°C.
3. Add 0.7 mL Phenol and slowly rotate at 4°C for 1 hr. - overnight.
4. Spin at high speed (15,000 g) for 2 minutes to separate phases. The hair and other debris should pellet to the bottom. Pipet the top (aqueous) phase to a fresh tube, taking care to avoid the bottom (organic) layer. Add 0.7 mL of a 1:1 (v:v) mix of chloroform and and phenol and rotate at 4°C for 1 hr.
5. Repeat step 3 with 100% chloroform.
6. Spin for 2 min, and transfer 0.5 mL aqueous phase to a fresh tube. Be careful not to transfer any chloroform at this step. Precipitate by adding 2 volumes (1 mL) of absolute ethanol and mix vigorously. Incubate at -20°C for at least 30 min. Samples may be stored in -20ºC in ethanol indefinitely.
7. Spin at 12,000 rpm for 5 min. Discard supernatant. Wash pellet by adding 1 mL of 70% ethanol, gently invert, and spin for 1 minute. Discard supernatant being taking care not to pour out the pellet which is likely to be loose adherent at this point. Repeat the 70% ethanol wash, spin and again discard the supernatant. Blot the excess ethanol carefully on a clean paper towel. Invert the tube on a paper towel to air dry the tube.
8. Resuspend pellet in 0.5 mL TE Use 1 µL for a PCR reaction, 0.25 mL for a Southern blot.

M. Fero. v.4

Materials

• RNA sample 30 µg Total or 2 µg mRNA
• Oligo dT (dT18VN) 5 µg/µL, 34.8 (µg/mL)/A260
• Superscript II, 5x 1st strand buffer (Invitrogen)
• 1 M DTT (Sigma, 43816, $16.80/10 mL)
• 50x dNTP Mix (dATP, dCTP, dGTP @25 mM, TTP=15mM)
• 10x AA-dUTP, 2 mM (MW = 523 g/mol), (Sigma A0410, $193.50/1mg - dissolve in 9.5 mL H2O)
• 1 N NaOH, 0.5 M EDTA, 2 M HEPES
• 0.2 M NaHCO3 (bicarbonate), 4 M NH2OH
• GFX spin columns
• QiaQuick columns
• PolyA (10 mg/mL) - Sterile filtered (0.45 µm)
• 20x SSC (175.5 mg/mL NaCl, 88.3 mg/mL NaCitrate)
• Cy3 and Cy5 Mono-reactive dye (Amersham PA23001 and PA25001):
  • To 1 tube dye add 36 µL anyhdrous DMSO and divide into 16x 2µL aliquots
  • Anhydrous DMSO: DMSO + Molecular Sieves (Sigma #M0133)

Notes

This procedure produces 1st strand cDNA with an anchored oligo dT primer and AA-UTP. It then couples couples amino reactive Cy-dyes to the AA-UTP. For spotted arrays control RNA should also be labeled using the opposing dye.

Procedure

A. 1st Strand Synthesis

1. In a 1.5 mL tube add:
   • RNA (30 µg total or 2 µg mRNA)
   • 1 µL (5 µg/µL) Oligo dT
   • q.s. 25 µL H2O
2. Incubate 70ºC, 10 min, then place on ice for 10 min.
3. Meanwhile, prepare RT Master Mix (per sample):
   • 2 µL Superscript II
   • 8 µL 5x 1st strand buffer
   • 0.4 µL 1 M DTT
   • 0.8 µL 50x dNTP mix
   • 4 µL 10x AA-dUTP (optional: spike with 1 µL 32P-dCTP)
4. Add 15 µL of RT Master Mix to each sample.
5. Incubate 42ºC, 2 hrs.

B. RNA Hydrolysis

1. To each sample add:
   • 12.5 1N NaOH
   • 5 µL 0.5 M EDTA
   • Incubate at 65ºC, 15 min.
   • Neutralize each sample with:
     • 20 µL 2M HEPES
     • O.K. to store o.n. at 4ºC.

C. Cleanup on GFX columns

1. Turn on Speed Vac refrigerators.
2. Add 0.5 mL GFX capture buffer. (optional: save 1 uL for 32P counts)
3. Load on GFX column. Spin 1 min. Discard flow-through.
4. Add 0.5 mL GFX wash buffer. Spin. Discard flow-through.
5. Elute in amber 1.5 mL tube:
   • Add 50 µL H2O. Incubate 1 min. Spin 1 min.
   • Repeat elution a 2nd time into the same tube. (Optional: save aliquot to measure 32P counts)
6. Dry in speed-vac.

D. Coupling to amino-reactive Cy dyes (e.g. Cy5 for sample, Cy3 for control)

1. Resuspend in 4.5 µL H2O
2. To a Cy-dye aliquot add: 2.25 µL 0.2 M NaHCO3
3. Quickly combine dye and DNA.
4. Incubate at r.t. in dark for 1 hr.
5. Quench by adding 4.5 µL 4 M NH2OH
6. Incubate at r.t. in dark for 15 min.

E. Cleanup in QiaQuick columns

1. Combine Cy5 and Cy3 samples (experiment and control).
2. Add: 70 µL H2O, 500 µL Buffer PB
3. Apply to a QiaQuick column. Spin 13K g for 30". Discard flow-through.
4. Add 750 µL Buffer PE. Spin. Discard flow through.
5. Repeat wash with PE.
6. Spin 1 min. to dry column.
7. Elute: Add 30 µL buffer EB, incubate 1 min. Spin 1 min. into a fresh amber tube.
8. Repeat elution 1x.
9. Speed-vac to dry down.

F. Preparation of Hybridization Solution

1. To each sample/control combination add:
   • 27.8 µL H2O
   • 5.4 µL 20x SSC
   • 2.8 µL polyA (10 mg/mL)
2. Load the DNA mixture on a 0.5 µm Millipore spin column
   • Spin at 12,000 g x5 min.
3. Store at -20ºC in the dark.

1. Prepare a 3 µL aliquot of each RNA sample at a concentration of 50 - 500 ng/uL in RNAse free H2O.
2. Submit to Genomics Core (cost = $4 / sample)

See Bioanalyzer manual for more details.

M. Fero 12/05

Materials

• MessageAmp II aRNA Amplication Kit (Cat #1751, Ambion)
• 100% Ethanol
• RNA samples (e.g. 1 µg RNA per sample)

Procedure

Reverse transcription

1. Verify that EtOH (24 mL) has been added to the Wash Buffer.
2. Turn on a 42ºC oven and set PCR machine to hold at 70ºC.
3. In an Eppendorf tube add:
   1. 1 µg Total RNA
   2. 1 µL T7 oligo(dT) primer
   3. q.s. to 12 µL with Nuclease-free H2O
4. Incubate 10 min. at 70ºC. Spin briefly to pull down any condensation. Place on ice.
5. Prepare RT-Master Mix (per sample):
   1. 2 µL 10x 1st strand buffer
   2. 4 µL dNTP mix
   3. 1 µL RNase inhibitor
   4. 1 µL ArrayScript
6. Add 8 µL RT-Master Mix to each RNA sample. Incubate at 42ºC for 2 hrs. Place on ice.

Second-strand cDNA synthesis

1. Set PCR machine to hold at 16ºC.
2. On ice prepare 2nd Strand Master Mix (per sample):
   1. 63 µL Nuclease-free H2O
   2. 10 µL 10x 2nd strand buffer
   3. 4 µL dNTP mix
   4. 2 µL DNA polymerase
   5. 1 µL RNase H
3. Vortex master mix briefly, pull down by spinning briefly and place on ice.
4. Add 80 µL 2nd Strand Master Mix to each RNA sample. Mix by pipetting, flicking and spin briefly to pull down.
5. Incubate RNA at 16ºC x 2 hrs on PCR machine. Do not close lid. Place on ice or freeze o.n.

cDNA purification

1. Preheat nuclease-free H2O to 55ºC.
2. Add 250 µL of cDNA Binding Buffer to each sample. Mix by pipetting and flicking, and spin briefly to pull down.
3. Load sample onto a cDNA Filter Cartridge in a wash tube. Spin 1 min. at 10,000 xg. Discard flow-through.
4. Add 500 µL Wash Buffer. Spin 1 min. at 10,000 xg. Discard flow-through.
5. Transfer cartridge to a cDNA Elution Tube.
6. Add 10 µL of 55ºC Nuclease Free H2O to the center of the cartridge. Incubate 2 min, then spin 1.5 min. at 10,000 xg.
7. Repeat elution in the same tube with another 10 µL of 55ºC H2O.
8. Store at -20ºC.

In vitro transcription (without biotinylation)

1. Heat oven to 37ºC.
2. Prepare IVT Master Mix (per sample):
   1. 4 µL T7 ATP Solution
   2. 4 µL T7 CTP Solution
   3. 4 µL T7 GTP Solution
   4. 4 µL T7 UTP Solution
   5. 4 µL T7 10x Reaction Buffer
   6. 4 µL T7 Enzyme Mix
3. Add 24 µL of IVT Master Mix to each sample. Incubate at 37ºC for 4hrs - 14 hrs.
4. Stop the reaction by adding 60 µL of Nuclease-free H2O. Vortex to mix. Store at -20ºC.

aRNA purification

1. Preaheat Nuclease-free H2O to 55ºC.
2. Label aRNA Filter Cartridges and place in aRNA Collection Tubes.
3. Add 350 µL of aRNA Binding Buffer to each aRNA sample.
4. Immediately add 250 µL absolute EtOH to each sample. Mix by pipetting and transfer to aRNA Filter cartridge.
5. Spin ~1 min. at 10,000 xg. Discard flow-through.
6. Add 650 µL Wash Buffer to each sample. Spin ~1 min. at 10,000 xg. Discard flow-through.
7. Transfer cartridges to fresh aRNA Collection Tubes.
8. Add 100 µL of 55ºC Nuclease-free H2O to each cartridge. Incubate 2 min. Spin ~1.5 min at 10,000 xg.
9. Store at -80ºC.

Matthew Fero, 4/5/06
(see Molecular Cloning 7.21-7.34)

Materials

DEPC H2O

DEPC 0.1% (v/v)
q.s. de-ioinized H2O
37ºC x1 hr, or r.t. overnight
Autoclave
(NaOAc, EDTA and ethidium bromide solutions should also be DEPC treated.
Tris has a reactive amine and can't be DEPC treated)

10x Formaldehyde gel loading buffer:

50% glycerol
10 mM EDTA, pH8
0.25% bromphenol blue
0.25% xylene cyanol

10x MOPS buffer (1L)

41.8 g MOPS (0.2M) in DEPC H2O, pH7
20 mL 1 M NaOAC (DEPC treated) (final 20 mM)
20 mL 0.5M EDTA, pH8 (DEPC treated) (final 10 mM)
q.s. 1L DEPC H2O
Sterile filter, store at 4ºC in dark. (Don't use if dark yellow)

1x Reaction buffer (per sample)

2 µL 10x MOPS buffer (final 1x)
4 µL formaldehyde (final 20%)
10 µL formamide (final 50%)
2 µL 0.2 mg/mL ethidium bromide, DEPC treated (final 10 µg/mL)

1.5% Agarose 2.2M formaldehyde gel

1.5 g agarose
72 mL H2O
Dissolve in microwave and then cool to 55ºC
In a fume hood add:
10 mL 10x MOPS buffer
18 mL de-ionized formaldehyde
Cast gel and wrap in Saran wrap until ready to use.

Note: Northern blots require formadehyde in the gel. For simple inspection, however, it is fine to use regular DNA-style TBE agarose gels. In either case, the RNA should initially be denatured (steps 2-3) and RNAse free reagents should be used.

Procedure
Add 2 µL RNA (up to 20 µg) to 18 µL 1x Reaction Buffer. Incubate at 55ºC x1 hr, or 85ºC x 10 min. Cool in ice and spin quickly to pull condensation down off of cap.
Add 2 µL 10x Formaldehyde gel loading buffer.
Load on agarose gel. Use formaldehyde agarose gels and 1x MOPS running buffer if doing a Northern or if accurate sizes are necessary. Otherwise 0.5x TBE gels are fine. Rinse gel box in DEPC H2O prior to use. Use RNA standards.
Run at 4-5 V/cm for 4 hrs. Place on Saran wrap prior to photographing

v.2 M. Fero 6/1/06

Procedure

1. Homogenize cells (10 million) or tissue (50-100 mg) in 1 mL TRIzol Reagent (e.g. scrape and pass through 30G needle, dounce homogenize and pass through needle, or use a homogenizer) and transfer to a 1.5 mL tube.
   Optional - Spin at 12k g for 10' at r.t. to pellet debris. Save pink supernatant in fresh tube.
2. Incubate 5' at r.t.
3. Add 0.2 mL CHCl3 (chloroform). Shake 15". Incubate 2-3' at r.t.
4. Spin 12K g 15', 2-8ºC.
5. Transfer clear (aqueous) phase to a fresh tube.
   Optional - Add 5 - 10 µg glycogen if < 10 mg of tissue was used at start.
6. Add 0.5 mL isopropanol, mix. Incubate at r.t. 10'. (Use 0.75 mL isopropanol for isolating miRNA)
7. Spin 12K g 15', 2-8ºC.
8. Discard supernatant. Add 1 mL 70% EtOH (with fresh with RNAse free H2O). Use 75% EtOH for miRNA. Vortex.
9. Spin 7500 g 5', 2-8ºC. Optional: Repeat 75% EtOH wash.
10. Discard supernatant. Air dry 5-10'.
11. Dissolve RNA in RNAse free H2O.

Fluorometer Quantitation

See Fluorometer protocol for more information.

1. Create a Ribogreen MasterMix: Turn on Fluorometer. Thaw 20x RNAse free TE, Ribogreen and RNA standard.
   • Master Mix (multiply by # samples + 2 standards)
   • 5 µL 20x T.E.
   • 0.5 µL RiboGreen (200x)
   • 92.5 µL DEPC treated H2O
2. Aliquot 98.5 µL master mix into 1.5 mL tubes.
3. Setup samples and controls:
   • Add 2 µL H2O into one tube as the "blank".
   • Add 2 µL of RNA standard into a tube as the "100 ng/uL" standard.
   • Dilute each sample 1/10 in DEPC H2O (1 µL + 9 µL water).
   • Add 2 µL dilute sample to the master mix aliquots.
4. Blank and then calibrate the fluorometer using the "blank" and "100 ng/µL" standard.
5. Measure each RNA sample. Multiply the results by 10x (dilution factor).

DNAse I Digestion

1. Create DNAse I buffer: MgCl2 and KCl should directly treated with 0.1% DEPC, incubated o.n. at room temp. and then autoclaved. Create 1M Tris pH8.4 by adding 4.03 g Tris base + 2.64 g Tris HCl in 50 mL DEPC treated H2O. (Tris buffers can't be treated with DEPC). Sterile filter.
2. Digest contaminating genomic DNA:
   • 100 µg RNA
   • 10 µL 10x DNAse I buffer (Tris pH8.4, 20 mM MgCl2, 500 mM KCl)
   • 5 µL DNAse I (RNAse free)
   • q.s. 100 µL with DEPC H2O
   • Incubate at room temp. x10 min. Immediately proceed to next step.

RNeasy Mini Prep

Procedure

(See RNeasy Mini Handbook for details)

1. Add 0.6 mL of RLT solution.
2. Transfer to a 1.5 mL tube.
3. Add 1 vol (0.6 mL) 70% EtOH. Mix by pipetting. Transfer 0.6 mL to a minicolumn in a collection tube.
4. Spin >10K rpm 15". Discard flow through. Return column to same tube.
5. Load remainder of lysate to column. Spin, discard flow through.
6. Wash by adding 0.7 mL RW1. Spin, discard flow through.
7. Wash by adding 0.5 mL RPE. Spin, discard flow through.
8. Repeat wash with 0.5 mL RPE, but spin for 2' to dry column.
   (Be sure no buffer is carried over on column tip.)
9. Elute by adding 50 µL RNAse-free H2O and transfer to a fresh tube. Spin >10K rpm 1'. Add another 50 µL H2O and spin again. Save pooled eluate at -20º or - 80ºC.

BioAnalyzer

1. Dilute ~50 ng of total RNA to 3µL with DEPC H2O, and submit to genomics lab.
2. Check account for results. Non degraded RNA will have a 28s/18s ratio = 2. The "concentration" result may be equated to the amount of RNA (ng) that was submitted. For more details on the interpretation of BioAnalyzer tracings see the section of the BioAnalyzer Nano 6000 kit manual that discusses assay results.

(M. Fero) 9/1/04

Protocol

1. To 1.5 mL eppendorf tubes add:
   • 200 µg of protein extract (see Western blot protocol for protein sample preps)
   • q.s. to 300 µL with RIPA (with protease and phosphatase inhibitors).
   • Primary antibody (amount determined emperically, approx 10x the concentration needed for westerns)
2. Vortex and incubate on ice for 30 min.
3. Meanwhile wash Protein A-Sepharose beads 2x in RIPA as follows:
   1. Cut off ends of pipetman tips
   2. Suspend beads and add beads to a 1.5 mL tube (40µL x # of reactions)
   3. Spin down beads, aspirate supernatant, resuspend in 1 mL of RIPA.
4. Resuspend beads in 1 vol. of RIPA (40µL x # of reactions)
5. Vortex beads before adding 30 µL to each reaction (a lot is lost on tips).
6. Rotate in cold room for 1 hr.
7. Spin and wash beads twice with RIPA (discard supernatant), and then once with Histone Wash Buffer.
8. Add 30 µL of Histone Assay solution. Incubate at 37°C for exactly 30 min. Mix by vortexing every 10 min.
9. Stop reaction by adding 15 µL of 4x sample buffer. Heat on 95°C block for 5 min. (Can be stored at 4°C overnight)
10. Spin down beads and load supernatant on a 12% SDS acrylamide gel. Run off the dye front (175 volts x 60 min. for a BioRad MiniProtean 3 gel)
11. Rinse gel in water. Stain with Coumassie x15 min. Destain for 1-2 hrs. (This will fix the proteins into the gel).
12. Wrap in Saran and expose to X-ray film at -80°C with intensifying screens. The optimal exposure time will depend on the amount of 32P used and the amount of kinase activity. Generally a few hours should be fine.

Solutions

Histone Wash Buffer

25 mM Tris HCl, pH7.5
70 mM NaCl
10 mM MgCl2
1 mM DTT

RIPA cell lysis buffer

10 mM NaPO4, pH7.2
0.3 M NaCl
0.1% SDS
1% NP40 (Nonidet P40)
1% DOC (sodium deoxycholate)
2 mM EDTA

Protease Inhibitors (add fresh to RIPA)

Leupeptin 5 mg/mL in PBS (1000-10,000x, aliquot and store -20°C), Roche # 11 017 101 001
Aprotinin 10 mg/mL in PBS (5000-160,000x, aliquot and store -20°C), Roche # 10 981 532 001
PMSF: (phenylmethylsulfonyl flouride, 100x) 0.2 M in ethanol. Store at 4°C.

100x Phosphatase inhibitor mix (store -20ºC and add fresh to RIPA)

100 mM NaF
50 mM NaVanadate
800 mM ß-glycerol phosphate

5x Sample Buffer - where should this link to?

S.R. — 1/2004

Procedure

1. Dilute p27 monoclonal antibody 1:1000 (v/v) in Carbonate Coating Buffer. Add 100 µl/well and incubate o/n @ 4°C or 1 hr @37°C.
2. Wash 3X with TBST (pour onto plate, empty into sink, hit onto towel 3x to clear wells).
3. Block plate by adding 270 µl/well of BSA/TBS solution.
4. Incubate @37°C for 1 hr.
5. Wash 3x withTBST.
6. Dilute samples and standards in assay buffer and add 100 µl/well (the highest concentration of standards is added to 200 µl then serially diluted in 100 µl assay buffer on the plate).
7. Incubate @ RT for 1 hr while shaking.
8. Meanwhile let TMB solution equilibrate to RT.
9. Wash plate 3x and add 100 µl/well rabbit polyclonal anti-p27 antibody diluted 1:1000 in assay buffer.
10. Incubate 1 hr @ RT with shaking.
11. Wash plate 3x and add 100 µl/well anti-rabbit HRP conjugated antibody diluted 1:1000 in assay buffer.
12. Incubate 1 hr @ RT w/shaking.
13. Wash 3x and add 100 µl/well TMB. Allow to develop w/o shaking for up to 15 min. @ RT.
14. Inactivate TMB with 1 N HCL and read @ A450 with TMB-S program in a plate reader.

Materials

TBST

TBS + 0.05% Tween 20

BSA/TBS

TBS + 2% w/v BSA

Antibodies

anti-p27 monoclonal antibody (BD Transduction Labs, 610241)

Rabbit anti-mouse p27 antibody (Fero lab)

TBM Eliza Color Substrate

3,3'5,5'-Tetramethyl-benzidine (TMB), 100 mL Sigma (T-0440)

Store at 4ºC

Plates

Nunc immuno-plate, C96 Maxisorp

M.Fero 12/2011
(Includes sample preparation, Bradford Assay, SDS-PAGE, semi-dry transfer, antibody staining and ECL development.)

Materials

See below.

Harvesting Cells

1. Prepare lysis buffer by adding fresh protease inhibitors (100x stocks listed below) to RIPA or TG buffer, plus 1/1000 vol. of PMSF (from a 200 mM stock).  Also add fresh phosphase inhibitors (from 100x stock) if kinase assays will be done. RIPA buffer is preferred, but it may be too harsh for some proteins. TG buffer, in contrast, may not lyse cells or nuclei as effectively, but it may be better for preserving cyclin D catalytic activity.
2. Scrape or trypsinize cells in culture. Spin and resuspend 107 cells in 150 µL lysis buffer. For tissues, dounce 100 mg. of tissue in 1 mL cell lysis buffer.
3. Sonicate sample on ice to fragment the genomic DNA. Be careful not to let samples overheat while sonicating (the tubes should remain cool to touch).
4. Spin at maximum speed, 4ºC to remove debris.  Transfer supernatant to a fresh tube.
5. To retain kinase activity it is important to not allow the samles to freeze solid.  Add 1 vol. glycerol (final = 50% v/v). The glycerol is viscous so, to facilitate pipetting, you should first cut the ends off of the pipetman tips with a pair of scissors.  Mix thoroughly by a combination of pipetting and vortexing.  Keep the extracts on ice when in use.  Otherwise store the extracts at -20ºC.  Ice crystals will form if the glycerol was not well mixed.
6. Prior to use, the protein content of samples should be assayed.

Protein Quantitation

Comments

The chief goal is to determine the relative protein abundance across samples to ensure equivalent loading of samples on the gel.  A second goal is to ensure that the absolute quantity of protein loaded is in a range that is sufficient for visualization but does not exceed the capacity of the gel.

If an abundant complex mixture is being assayed (e.g. cell or tissue extracts) then an A280 measurement may suffice.  However, A280 measurements are dependent on the amino acid composition of the sample, chiefly tryptophan and tyrosine, so A280 readings may not be appropriate for adjusting the concentrations of two different purified proteins or two different tissue types. A280 readings are sensitive to the presence of contaminating nucleic acids (DNA and RNA).

The Bradford assay is relatively easy, sensitive, and less dependent on amino acid composition. It has a limited linear range, so it should be repeated on samples that give high readings and are subsequently diluted. It is recommended to measure all samples, that will be run on a single gel, together in a single Bradford assay, since there may be day to day variability in the results.

Bradford assay:    

• Dilute Bradford reagent to 1/5x by adding 1 vol. reagent + 4 vol. H2O.
• Aliquot 0.8 mL diluted Bradford reagent into 1.5 mL tubes
• Add 2 µL of protein extract and mix by vortexing.  Incubate 2-5 min. at room temp.
• As a negative control add 2 µL of your protein lysis buffer to a separate 0.8 mL aliquot of Bradford reagent.
• Zero the spectrophotometer at 600 nm using the negative control.
• Measure A600 for each sample in the same order that they were prepared (since the Bradford reagent absorbance will gradually increase over time).
• To calculate the approximate protein concentration (µg/µL) multiply A600 x 10.  In order to ensure relative precision it is best if all of the extracts to be run on a single gel are quantified simultaneously.  If necessary more accurate values may be obtained by adjusting the results according to a standard curve of a known protein.

   SDS PAGE (Volumes used for 1 mm BioRad Mini-Protean gel system)

(Tris buffers must be made from Tris-base, and are pH'd with conc. HCl.  Store acrylamide, 10% APS, and TEMED at 4ºC.)

7. Use 4 mL of resolving buffer for a 1mm MiniProtean gel. Gently overlay resolving buffer with ethanol to shield the buffer from air which will inhibit polymerization. Rinse off ethanol with water when gel has polymerized, and drain out the water.
8. Overlay with stack gel and place a 10 or 15-well comb.
9. Prepare 10 - 50 µg protein in 15 - 25 µL of lysis buffer per lane (the volume depends on comb size). Add 1/4 vol of 5x SDS loading buffer to each sample. Heat on a 95°C block x 3 min prior to loading, then hold on ice.
10. Load samples into wells, along with lanes dedicated to a prestained MW marker and (+) and (-) controls.
11. Run at 200 v. for 1 hr or until dye front runs off the bottom of the gel.  Thicker gels (1.5 mm) will run hotter and should be nearly submerged in running buffer or run at lower voltages.

Electrotransfer (using Ellard Instruments HEB 2020 semi-dry blotter):=   

12. Cut 15 Whatman 3M filter sheets to 5.5 x 8.5 cm and one PDVF membrane by the same dimensions.
13. Soak filter paper in buffers A (6 sheets), B (3 sheets), and C (6 sheets).
14. Wet PDVF membrane in methanol. Hydrate in H2O,then equilibrate in buffer B.
15. Separate glass plates. Rinse gel in H2O. Discard stack. Create transfer sandwich on Saran wrap. (Soak filters in the appropriate solutions for 2 min. and squeeze out the extra solution):
    Sandwich from bottom to top:
    Saran wrap (on lab bench)
    Buffer A filters (6 sheets)
    Gel
    PVDF (prewetted in B).
    Buffer B filters (3 sheets)
    Buffer C filters (6 sheets)
16. Invert this sandwich onto the base (+) eletrode of the transfer apparatus. Remove saran wrap. Place (-) electrode on top of sandwich.
17. Transfer for 1 hr. at 40 mA per blot for 1 mm thick gels.  (The protein will migrate out of the gel towards the (+) electrode and will stick to the PVDF membrane).
    Note: The necessary current is a function of the surface area of the gel sandwich so the mA must be increased proportional to the number of gels being run, e.g. use 160 mA if blotting for gels simultaneously.  The running time should be proportional to the thickness of the gel, so this may be reduced to 45 min. for 0.75 mm gels, or increased to 90 min. for 1.5 mm gels. High MW proteins (> 200 kD), or high percentage acrylamide gels may require longer transfer times.

Antibody Staining  

18. When transfer is complete:  Remove filter paper sandwich from electro-blotter apparatus. Mark MW bands with ink from a ball point pen (Papermate ink won't wash out). For orientation, nick the corner above MW markers.  Meanwhile, stain the residual proteins in the gel, with Coumassie blue (see below).
19. Stain the PDVF membrane with antibody as follows (after each step rinse well in several changes of TNT x 10 min):

• 5% Milk in 0.5% TNT (2.5 gm milk in 50 mL TNT) x 30-60 min. Rinse in TNT.
• Primary antibody, 8 mL x 1 hr.(most are 1/1000 in 0.5% TNT). Rinse in TNT.
• Secondary antibody 8 mL (1:10,000 HRP-anti-rabbit IgG) Rinse in well in TNT
• Add ECL mix (1:1 mix of Amersham reagents 1 and 2) and immediately expose to film x 1 min. Alternatively, if an appropriate fluorescently-labeled secondary antibody was used, then the blot may be scanned on an Odyssey imager (Li-Cor).

Coumassie Blue Staining

Note: Following electrotransfer a small amount of protein (~10% of the total) will remain in the gel, assuming a 1mm gel was blotted for the time listed above. If a thinner gel is used then only a small amount of high MW material may remain or else the electrotransfer time may be proportionally reduced. Having a small amount of residual protein in the gel is convenient because it can be post-stained with Coumassie blue in order to document consistent protein loading across samples. Alternatively, membranes may be stained with Ponceau S solution (Sigma) for the same purpose.

20. When the gel is removed from the transfer stack (in step 13) rinse it briefly in H2O and place it in a staining tray. Gently agigate the gel in Coumassie Stain for 30 min.
21. Rinse in a small volume of Destain and then cover in Destain in a tray and gently aggitate for one hour. Repeat 1-2 times as necessary. Keep a balled up Kimwipe in the tray while destaining in order to absorb stain particles and maintain clarity of the destain solution.
22. Photograph the gel on a white background for documentation.
23. After use return the Coumassie stain to a glass bottle as it can be reused multiple times. The Destain solutions should be discarded with organic wastes.

Interpretation of Results

Western blots are usually not quantified and thus are limited to qualitative interpretations, e.g. "The amount of protein X is higher in sample 1 than in sample 2", or "The levels of protein Y is unchanged across the samples". Still, some investigators choose to immunostain blots a 2nd time with an antibody against a protein, such as actin or tubulin, which are expressed at stable levels, as a "protein loading control". This gives visual reassuance that the changes seen for the protein of interest was not due to technical problems with protein loading. Regardless, of whether such a control is used, researchers should establish that both the experimental protein (and the loading control) are expressed within the "linear range" of the assay. For example, ECL may exhibit a threshold effect wherein the reduction of the target protein below a certain level is associated with a disappearance of a band rather than a reduction of its intensity.  The detection system may also become "saturated" for proteins with high expression levels, such that significant differences in levels will appear to be the same.  This is often the case for actin loading controls if a high concentration of antibody is used.

Quantitation

Western blots can be quantified with a reasonable level of accuracy if one is careful about the technical setup. A traditional way to do this is to scan films and perform densitometry. Fluorescent antibodies can also be used, in conjunction with an Odyssey imager, and exhibit a greater dynamic range and accuracy than densitometry. In either case, a standard curve comprised of two-fold serial dilutions of a positive control should be run in parallel to verify the linearity of the assay and to quantify the results. Minor differences in protein loading may be normalized by the differences seen in the actin or tubulin internal control, assuming that this too exhibits a linear relationship in a standard curve.

MATERIALS

Protease Inhibitors

Leupeptin 10 mg/mL (1000x) Store at -20°C.
Aprotinin 10 mg/mL (1000x) Store at -20°C.
PMSF: Phenylmethylsulfonyl flouride, 200mM in ethanol (100x) . Store at 4°C.

100x Phosphatase inhibitors (for kinase assays)

100 mM NaF
50 mM NaVanadate
800 mM ß-glycerol phosphate

Western Transfer Solutions

Solution A: 25 mM Tris Base, 20% v/v isopropanol, 40 mM e-aminocaproic acid.
Solution B: 25 mM Tris Base, 20% v/v isopropanol.
Solution C: 250 mM Tris Base, 20% v/v isopropanol.

Coomassie Stain and Destain

Coomassie Stain: 0.25% w/v brilliant blue (Sigma B-0770), 50% v/v methanol, 7.5% v/v glacial acetic acid. Filter through Whatman #1.
Destain: 10% (v/v) methanol, 10% (v/v) glacial acetic acid.

0.5% TNT: 0.5% Tween-20, 0.15 M NaCl, 25 mM Tris pH 7.4.

Non-fat dried milk.

M.Fero — 11/94

Procedure

1. Prepare serial 10-fold dilutions of transformed bacteria in LB and spread 100 µL onto LB/Amp plates, as described in the Electroporation protocol. Incubate at 37°C overnight. Also streak out positive and negative control bacteria on a separate plate.
2. Select plates that have an optimal density of bacteria (i.e. 2 mm spacing between colonies). Put selected plates at 4°C for one hour.
3. Lay 83 mm circular nylon membrane on surface of agar plate until it becomes thoroughly wetted. Poke 3 widely spaced wholes in an asymmetrical pattern through the filters and the agar near the edge of the dish with a 18G or 21G needle for future orientation (see Figure 1).
4. Place Saran wrap on benchtop. Spot out 0.5 mL puddles of denaturing solution (x1) and neutralizing solution (x2) for each filter at 6" intervals on Saran wrap.
5. Using forceps, carefully peel nylon membrane from agar surface and place it colony side up onto a puddle of denaturing solution. The colonies should stick to the membrane and not to the plate. Incubate for 5 min. Blot it briefly on a paper towel (colony side up) and then transfer it to the first puddle of neutralizing solution for 5 minutes. Repeat this with a second puddle of neutralizing solution.
6. Cross link the DNA under a UV light (Stratalinker).
7. Rinse the filters in 2X SSC + 0.1% SDS on a rotator.
8. Lay the filter colony side down onto a paper towel. Lay a second towel on top and gently press on the filter to blot away most of the lysed bacterial protein. Be careful not to smear the colonies. They can be kept moist or dry until hybridization.
9. Regrow colonies on the plates by returning them to 37°C for 4 hrs and then store them at 4°C.
10. Hybridize the filters and wash them as usual. Monitor the washes with a Geiger counter to be sure that most of the counts have been washed off. If an oligo is used as a probe then the hybridization should be done in the refrigerator without formamide and the washes should be done at low stringency (1x SSC) at room temperature.
11. Dry the filters and lay them out on an old piece of film. Place fluorescent markers at the corners of the film. Wrap the film in Saran wrap to hold the filters in place. Expose to film at –70°C for 4 hrs to overnight.
12. Develop and dry the autoradiograph. Line up the autorad with the filters such that the fluorescent markers line up precisely. Dot the locations of the filters pinholes on the autorad with a Sharpie. Place the autorad on a light box (see Figure 2). Place the bacterial dishes on the autoroad lining up the agar pinholes directly over the ink marks on the film. Pick colonies that correspond to the autorad spots.

Materials

LB Amp plates
Positive and negative control bacteria
Saran wrap
Denaturing solution (0.5 M NaOH + 1 M NaCl)
Neutralizing solution (1 M Tris pH 7.4, 1 M NaCl)
2X SSC, 0.1% SDS
83 mm circular nylon membranes (Hybond N)
Stratalinker UV box
Radiolabled probe
Hybridization Solution (e.g. Stark’s)
Autoradiograph film and cassettes

M.Fero 11/30/2011   (links to be added to other protocols once they are entered)

Large Scale Preps

(See Large scale plsasmid prep protocol for more details)

1. Cultures: Inoculate a 5 mL LB/Amp (50 - 100 µg/mL) culture in early a.m. with a single colony. Use all 5 mL to inoculate a 500 mL LB/Amp culture in the evening. Alternatively the 5 mL culture can also be set up as an overnight culture.
2. Save 1 mL for a glycerol stock if necessary (see step 6c, below).   Prepare remainder according to alkaline lysis PEG or CsCl/Qiagen protocol.
3. Measure and record A260/A280.  Perform diagnostic digests on 0.1 µg of DNA, e.g. digests which will compare uncut, linearized, cut out insert, and cleaved insert.  Alternatively, sequence the insert using vector primers which flank the cloninig site (e.g. T3, T7, or M13 primers)
4. For lab stocks record on each tube: Plasmid name, plasmid ID number (red ink), your initials, date of plasmid prep or lab book page number, DNA concentration.
5. Consider reprepping the DNA if the A260/A280 is less than 1.88, if their is any visible protein in the wells of the test gel.
6. For newly acquired or constructed plasmids:
   1. Draw plasmid map using GCK software and print a copy for the lab notebook.  Also record the plasmid name and location in the MPD database.
   2. Store the plasmid at -20°C. In the lab plasmid box.
   3. For commonly prepped or particularly valuable plasmids store a glycerol stock as follows: Add 200 µL of glycerol to 1 mL of an overnight culture of the bacteria and mix well. Store at -80°C and record the plasmid name, I.D. number, date and the strain of bacteria in the MPD's Freezer and Plasmid databases. This will speed up future plasmid preps and ensure that not all of the plasmid is used up.

Test Digests

A typical restricion digest consists of:  0.1 µg of DNA + 1 µL of enzyme + 2 µL of 10x buffer (consult Roche or NEB list) and q.s. to 20 µL with H2O.  Note that some NEB buffers also call for the addition of BSA.  Digest for 2 hrs at the temperature recommended for the enzyme (usually 37ºC). (Longer digests are not advisable for miniprep DNA because it will increase the likelihood of degragradation of the DNA by exonucleases).

Test Gels

1. Add 1/10 vol. of tracking dye to 0.1 µg DNA per lane. Use more DNA only if you need to visualize both very small and large fragments. Do not overload the wells as this will cause bands to smear. Add 4 µL of 10 mg/mL ethidium bromide per 100 mL of 0.5x TBE with the following concentrations of agarose:

2. Run the medium sized gel boxes at 100 v. for 90 min - 2 hrs.  Run the small boxes at 80 v. To resolve very large fragments (>10 kB) load an equivalent of 20 ng per large fragment. Run at a reduced voltage (e.g. 50 v.) for 4 - 6 hours.

Preparative Digests

Note:  If a digest will produce only a single fragment (e.g. a vector being cut with a single enzyme, or PCR product cut at their ends) then the DNA should be digested, phenol extracted and purified by EtOH precipitation.  If the digest produces a product which must be purified away from a second fragment (e.g. insert removed from a vector) then the DNA should be digested, run on a prep gel, and then EtOH precipitated.  Agarose gel electrophoresis has the disadvantage of reducing the cloning efficiency of DNA fragments.

1. Calculate the optimal reaction conditions to obtain 5 µg of DNA of the fragment of interest:
   • Amount of DNA = 5 µg x (size of fragment)/ (total plasmid size).
   • Use 10 units of enzyme per µg of plasmid DNA.
2. To ensure that the DNA is cut to completion it should be cut under dilute conditions:
   • 5 µg of vector DNA + 50 U. enzyme + 50 µL 10x buffer, q.s. to  0.5 mL with H2O.
3. Digest for 90 min. at the optimal temp (usually 37ºC). Add more enzyme (again 10 U/µg) for another 90 minutes.
4. Pour two gels: A mini-test gel and a mid-sized prep gel. Use 10-tooth combs for the preparative gel.  When the DNA is half way digested remove 0.1 µg for a test gel with 0.1 µg of uncut DNA.
5. If the test gel shows >90% digestion load 1/2 of the preparative sample into 3 - 5 lanes of the preparative gel. Run at <75 volts to mimimize smearing of the bands.
6. Photograph the prep gel when it is finished running. Indicate on the photo which band was cut out. If there are multiple bands photograph the gel after the band is removed.
7. Elute the DNA from the agarose gel.  There are several protocols for doing this including elution onto glass fiber filters, elution into dialysis tubing, and use of an elution trap.  (See Moleclular Cloning for more details).  Precipitate the DNA with NaOAc and EtOH, wash with 70% EtOH, and resuspend in TE or H2O.
8. Quantify and record the estimated DNA concentration of the isolated gel fragment on a test gel with Lambda + HindIII size marker.  Calculate the molar ratios of vector to insert.

DNA Ligation

Typical reaction conditions for directional cloning ligation:
20 ng of vector
3-fold molar excess of insert
2 µL of ligase
20 µL reaction volume.
Ligate at 15°C for 2 hours to overnight.

For large or problematic cloning steps it may be informative to double the recipe so 1/2 of the sample can be run on a test gel and 1/2 can be used for bacterial transformation.  In this case vector only and insert only ligation controls should also be performed in parallel (see Bacterial transformation, 4a, below).  If the appropriate ligation products are not present on the test gel then the DNA may need to be repurified or the ligation conditions may need to be optimized (see Ligation Optimization protocol).

Bacterial Transformation

1. Use highly competent cells for blunt ended cloning or very large constructs. Transform only 1/2 of the DNA into bacteria and incubate in 1 mL of LB for 30-60 min. (Save the remaining 1/2 of the ligation mixture to run on a gel to check the ligation competence of your fragments).
2. Blue/White selection: Evenly spread 120 µL X-gal/IPTG across plates > 1 hour before use.  See Blue-White selection protocol for more details.
3. Plate 100 µL of bacteria at different dilutions in LB: 1/100, 1/10, 1x, and 10x (for the latter spin 1 mL of culture and resuspend in 0.1 mL LB).
4. Plate 100 µL of controls:
   1. 1 µL unligated vector fragment,
   2. 1 µL unligated insert fragment,
   3. 0.1 ng of uncut pBS. Record the plating efficiency (# colonies pBS x 105 /µg pBS)
   4. Pick individual colonies and restreak them onto LB/Amp plates and simultaneously innoculate LB/Amp 5 mL broth cultures for minipreps.

DNA Minipreps

1. Grow 28 overnight cultures each in 5 mL of LB/Amp broth.
2. Miniprep 1.5 mL of bacteria with an alkaline lysis protocol with RNAse and 1 phenol or chloroform extraction followed by 1 ethanol precipitation with 2x 70% EtOH washes. Alternatively, use a Qiagen mini-prep kit.
3. Resuspend miniprep in 50 µL of T.E. Use 3 µL per diagnostic digest. Pour 2 medium test gels each with two 15-tooth combs. Run test gels with both uncut and cut DNA.  If miniprep digests fail to find inserts you may use the perform colony hybridization on bacteria plates with 100 - 500 colonies.
4. Confirm orientation and secondary cut sites of insert of the relevant minipreps. Sequence all PCR cloned and blunt ligated fragments from both directions to confirm that no mutations have been introduced.  For PCR cloned fragments it is important to sequence the entire insert region to rule out point mutations.
5. Repeat digests with more enzyme if there is evidence of partial digestion. The presence or RNA (usually seen at the bottom of the gel) may also inhibit DNA digestion.  It may be necessary to repeat minipreps if the DNA is significantly smeary, degraded, contaminated with RNA, or contaminated by protein (material stuck in the wells of the gel)

M. Fero — 12/29/00

1. Grow 250 ­ 500 mL of bacteria overnight in LB with 50 µg/mL of ampicillin.
2. Transfer culture to Nalgene bottles. Spin in SGA rotor at 6000 rpm, at 4°C for 10 min.
3. Discard supernatant. Keep everything on ice. Resuspend bacterial pellets by adding 15 mL of water and split into 2 plastic Sorval tubes (max capacity = 40 mL/tube)
4. Add 17.5 mL of fresh Solution II to each tube. Mix by inversion. Keep on ice for 10 min.
5. Add 13 mL of Solution III. Mix by sharp inversion. Ice for 20 min.
6. Spin for 20 min. at 8,000 rpm (12,000g) in a JA-10 rotor.
7. Transfer supernatant to a 50 mL conical tubes by filtering through Kimwipes, use glass pipettes and avoid white cheesy gunk.
   • Split evenly into 4 glass Sorval tubes (max 20 mL/tube) Add an equal volume of isopropanol to each tube. Mix.
   • Spin 15 min. at 8,000 rpm (12,000g) in a JA-10 rotor.
8. Discard supernatant. Air dry pellet.
   • Resuspend pellets in a total of 4 mL of T.E. with 100 µg/mL of boiled RNAseA and transfer to 15 mL conical polypropylene tubes. Incubate at 25°C for 30 min. Add 0.5 mL of 5 M NaCl and 100 µL of 10% SDS, mix.
9. Extract 2x with an equal volume of phenol. Extract 1x with CHCl3 (with 1/25 v/v isoamyl alcohol. Repeat the extractions if there is still gunk left in the aqueous phase.
10. Transfer aqueous phase to 15 mL Corex tubes, add 9.4 mL of cold ethanol, mix. (It's OK to store the samples in the freezer at this point)
    • Centrifuge at 10,000 rpm, 10 min, 4°C.
11. Resuspend pellet in 500 µL of TE. Add 120 µL 5M NaCl, mix. Add 400 µL of 20% PEG 8000. Incubate on ice for 20 min. Pellet at 14,000 rpm, 10 min.
12. Discard supernatant. Pulse spin and discard remainder of supernatant.
    • Resuspend in 200 µL of TE. Add 200 µL 5M NH4OAC. Ice 20 min. Spin at 14,000 rpm. Save supernatant.
    • Add 1 mL of cold EtOH. Incubate at ­70°C for 5 ­ 10 min. or longer at ­20°C.
    • Rinse 2x with 70% EtOH. Air dry pellet.
13. Resuspend in 500 µL of TE. Check A260 and A280 and inspect on agarose gel.
14. Store glycerol stocks of the bacteria and make plasmid maps if they are not already done.

Solutions

• Solution II: 0.2M NaOH, 1% SDS. (Make fresh. For 20 mL add 0.4 mL 5M NaOH, plus 2 mL 10%SDS)
• Solution III: 3M KOAc, pH4.8. (60 mL KOAc, 11.5 mL glacial HOAc, 28.5 mL water).
• Isopropanol, Ethanol, 70% ethanol
• RNAseA: 10 mg/mL in water (boil for 20 min. Spin to remove debris. Freeze in aliquots)
• Phenol, equilibrated with TE, pH8
• Chloroform (+ Isoamyl alcohol 1/25 v/v).
• 20% PEG 8000

The following protocol can be used to optimize ligation conditions for difficult to clone (e.g. very large) fragments. The principle is to independently characterize the ligation kinetics of the vector and insert DNA fragments and then to combine them in optimal ratios. The final ligation is also performed in two stages to optimize the proportion of vector:insert heterodimers followed by a shift to low concentration to optimize recircularization. Reactions are performed in sufficient quantities to permit analysis on agarose gels which allows reaction optimization, and identifies problematic substrates with unligatable ends or contamination with ligase inhibitors.

Materials

T4 DNA ligase (and buffers from both Invitrogen and Roche).
0.1 M EDTA
3M NaOAc
70% EtOH
Glycogen 20 mg/mL (Roche)
TBE or TAE agarose gel
LB-Amp plates

Procedure

1. Gel purify DNA restriction fragments of interest for both the vector and insert. Quantify the isolated fragments on a test gel with a DNA concentration standard.
2. Optimize ligase concentration with fixed amount of Insert DNA:
   Set up the first test ligation with serial dilutions of ligase to find the optimal enzyme concentration: Use 20 ng of insert DNA, 3 µL 5x Invitrogen ligation buffer, ligase and q.s. to 15 µL with water. Use 1 µL of enzyme in the first tube and then perform 10-fold serial dilutions of enzyme in 5 tubes. Incubate at room temperature for 15 minutes. Stop the reaction by adding 1/10 vol. of 0.1M EDTA and heat to 65°C for 5 min.
3. Run a TBE agarose test gel with 0.4 µg/mL ethidium bromide to resolve the ligated multimers. Use unligated DNA as a size control. Bands which run smaller than the unligated DNA are circular forms. Choose an enzyme concentration which gives maximal amount of dimers. If circular forms predominate then the DNA concentration was not high enough or there was no PEG in the ligase buffer.
4. Optimize Vector DNA concentration:
   Set up the second test ligation with serial dilutions of vector DNA to find the concentration which is optimal with the chosen enzyme concentration. Set it up as in step 2 except use a fixed concentration of ligase and instead titrate the vector DNA concentration (perform 2 or 4-fold dilutions). Run a test gel again and choose the DNA concentration which gives you optimal dimer formation. Alternatively, you can arbitrarily choose a vector DNA concentration which is 3-4 fold less (on a molar basis) than was used for the insert and skip to step 5.
5. Ligate Vector and Insert to form a heterodimer:
   Set up the final ligation as a two-step reaction: Use 20 ng of insert DNA, the optimal dilution of enzyme, and the optimized quantity of vector DNA. Ligate at room temperature for 15 minutes at room temperature. Remove and heat inactivate activate a 5 µL aliquot for a test gel. It is a good idea to also ligate vector only and insert only under the same condtions.
6. Recircularize heterodimers:
   Dilute the remainder of the ligation mix to a total volume of 250 µL with water, 2 µL of enzyme and 23 µL of 10x Roche ligation buffer (which does not contain PEG). Incubate at room temp. for 4 hours or overnight at 15°C.
7. Ethanol precipitate by adding 25 µL of 3M NaOAc, 1µL 20 mg/mL glycogen and 550 µL of ethanol at -20°C for 30 minutes. Spin for 10 minutes, and wash the pellet 2x with 70% ethanol. Resuspend in 5-10 µL of water and transform half of this into bacteria, and run the remainder on a test gel with the aliquots that were previously saved. On the test gel you should see both multimeric forms and circular forms. Occasionally, the addition of vector DNA to insert (or vice versa) will cause an inhibition of ligation. This is often due to contaminants from agarose gel purification. Agarose gel purification of vector DNA can often be avoided altogether by taking care to cut the vector DNA to completion (e.g. cut the DNA twice with phenol extraction and EtOH purification after each digestion).

Because optimized ligation reactions may produce a large number of transformants it is important to perform 10-fold serial dilutions of the transformed bacteria while plating on to LB/Amp plates. If plasmid miniprep digests fail to identify the appropritate clone then plates the plates may be used for screening by colony hybridization.

M.Fero — 10/26/99

Materials

Solution II: 0.2N NaOH/1% SDS
Solution III: 3 M KOAc, pH4.8
RNAseA (DNAse free) 10 µg/mL
Chloroform/Isoamyl alcohol (1/25 v/v)
Isopropanol
70 % ethanol
13% PEG8000

Procedure

1. Culture 5 mL of bacteria o.n. in LB + 100 mg/mL ampicillin.
2. Spin down 1.5 mL of each culture (5,000 rpm x 3 min.) Discard supernatant. Add additional 1.5 mL of cell suspension and spin a second time.
3. Resuspend pellet in 100 µL of H2O.
4. Add 300 µL of Solution II. Mix, incubate on ice for 5 min.
5. Add 300 µL of Solution III. Mix, incubate on ice for 5 min.
6. Spin 5 min at maximum speed, 4°C. Save supernatant in a fresh tube.
7. Add 10 µL of RNAseA, incubate at 37°C x5 min.
8. Extract with 1 volume of chloroform x1, spin 2 min. Transfer aqueous phase to a new tube.
9. Add 1 volume of isopropanol, mix. Spin at max. speed x 2 min.
10. Rinse with 70% ethanol. Dry pellet. Resuspend in 80 µL of T.E.

Optional (for DNA sequencing)

11. Add 20 µL of 4 M NaCl, mix. Add 100 µL of 13% PEG, mix. Incubate on ice x20 min.
12. Spin at max speed, 4°C x15 min. Rinse pellet with 70% ethanol. Dry pellet.
13. Resuspend in 50 µL H2O.

8/17/2011 Shyamala Mohan
Protocol adapted from Molecular cloning 3rd edition, Joseph Sambrook and David W. Russell

Electrocompetent cells are prepared by growing cultures to mid log phase, washing the bacteria at low temperature and resuspending them in a solution of low ionic strength containing glycerol. The starting material can be a vial of compentent cells from a prior preparation. If there is any uncertainty about the purity of the cells they should first be cultured by streaking onto LB and LB + Amp plates. Untransformed competent cells should only grow on LB plates. Single colonies can then be picked and grown in LB media to generate a culture. The final product of this preparation are aliquots of bacteria that can then be be used for DNA transformation using electroporation.

Materials

DH5α E. coli, or comparable strain
LB culture medium
Sterile 10% glycerol in dionized water (40 mL of glycerol in 360 mL water)
Ice water bath
LB agar plates
LB + Amp agar plates
Sterile centrifuge bottles (250 or 500 mL)
Spectrometer and cuvettes
Dry ice/ethanol bath

Procedure

1. Thaw 50 µl of frozen DH5α competent cells and inoculate it in 500 mL of LB media. Inoculate the flask at 37ºC with agitation. Measure the absorbance (A600) of the growing bacterial culture every 20 minutes. To estimate the finish time, wait until the cultures reach exponential growth and then graph the A600 on semi-log plot (time vs log A600) to estimate the time for an A600 of 0.4.
2. In preparation for the next step, place the centrifuge bottles in ice-water bath. Place the 400 mL of 10% glycerol and 500 mL water in ice bath. Turn on the Sorvall centrifuge and cool the chamber to 4ºC with the rotor in place. Also precool a table-top centrifuge. These steps are important to keep the bacteria cold.
3. Bacteria are most competent in mid log-phase (A600 = 0.4-0.5). Measure the A600 by removing 0.5 mL into a plastic cuvette. Record the absorbance at 600 nm. When the A600 > 0.4, transfer the flasks to an ice water bath for 15-30 minutes. Swirl the culture occasionally to ensure that cooling occurs evenly.
4. Transfer the cultures to ice cold centrifuge bottles. Harvest the cells by centrifugation at 1000 g (~2500 rpm) for 15 minutes at 4ºC. Decant the supernatant and resuspend the cell pellet in 500 mL of ice cold pure H2O.
5. Repeat the spin and wash of the bacteria, but this time resuspend the bacteria pellet in 250 mL of ice cold 10% glycerol.
6. Repeat the spin and wash of the bacteria, but this time resuspend the bacteria pellet in 10 mL of ice cold 10% glycerol. Meanwhile place 15 mL conical tubes on ice.
7. Harvest cells by centrifugation at 1000 g for 20 minutes at 4ºC in a table top centrifuge. Carefully decant the supernatant and resuspend the pellet in 1 mL of 10% ice cold glycerol. Meanwhile, place 1.5 mL Eppendorf tubes on ice.
8. For storage, dispense 50 µL aliquots of the cell suspension into sterile ice-cold 1.5 mL microcentrifuge tubes, drop into a bath of dry ice with ethanol (to freeze the competent cells immediately) and transfer to a -80ºC freezer. Record the location of the cells in the MPD.
9. On the following day test the competency of the cells by transforming aliquots with successive dilutions of pBS plasmid vector. Count the number of colonies on a plate and calculate the number of colonies per µg of DNA transformed. Highly competent cells should yield 108 to 109 colonies per µg of DNA transformed. High levels of competency are needed for difficult cloning procedures, but lower competency cells can still be used for maintaining plasmid stocks.

M. Fero — 4/04

Procedure

1. Desalt DNA template by EtOH precipitation in NaOAc followed by at least 2x washes with 70% EtOH. Resuspend in 5 - 15 µL of sterile H2O.
2. Rinse cuvettes (if they have been used before) 5x with deionied H2O, and place them on ice. This is sufficient to avoid background growth in most cases.
3. Set a BioRad MicroPulser to "Ec1" for 1 mm cuvettes, or "Ec 2" for 2 mm cuvettes.
4. Electroporate the DNA into the bacteria:
   1. Add 5 µL of DNA to 50 µL of bacteria and mix by pipetting.
   2. Transfer bacteria/DNA mix to a cold cuvette, and immediately pulse in the electroporator.
   3. Quickly add 1 mL of r.t. L.B. to the bacteria in the cuvette. Use a sterile Pasteur pipette to transfer the suspension to a bacteria tube.
   4. Repeat for additional DNA samples and control DNA.
5. Rotate at 37ºC x30 min. to allow the bacteria to recover.
6. Meanwhile plate LB/Amp plates with IPTG + X-gal if blue/white selection of colonies will be performed, as follows:
   1. Mix 100 µL X-gal + 20 µL of IPTG per plate.
   2. Spread X-gal/IPTG mix across surface of plate with a sterile glass spreader and a plate spinner.
   3. Allow the mix to infiltrate the media for 20 min.
7. Perform two 10-fold serial dilutions of the LB/bacteria suspension into fresh sterile L.B. Concentrate the remaining bacteria by spinning 1 min. at 5k rpm in a sterile microcentrifuge tube, and then resuspend pellet in 0.1 mL of L.B.
8. Plate 100 µL of the various concentrations of bacteria onto LB/Amp (+ X-gal/IPTG as necessary), and spread with a sterile spreader and plate spinner to evenly coat the plate. Record the DNA construct and the Bacteria Dilution Factor (9x, 1x, 1/10x, 1/100x) on each plate.
9. Incubate the plates inverted at 37ºC o.n. They may need to be placed in a container if the humidity of the incubator is so low that it causes the agar to dry out.
10. Remove the plates when the colonies are ~1mm in diameter. The color development on X-gal treated plates will continue to occur after the plates are removed from the incubator.
11. Pick white colonies (also with no central blue coloration) and restreak on L.B./Amp plates to ensure that pure clones are obtained before performing plasmid preps.
12. Store at 4ºC for at least 1 hr. (to firm up the agar) if colony hybridiation will be performed. Seal edges of plates with parafilm for storage up to 1 week.
13. Count the numbers of colonies on the plate with control DNA to determine the efficiency of competent cells:
14. Efficiency (Transformants/µg) = colonies/plate x (Bacterial dilution factor) x 20,000
15. Highly competent cells should have ~50 colonies on the plate with a 1/10 dilution of the control DNA's bacteria suspension.

Materials

Control DNA (e.g. pBS) diluted to 1 pg/µL in sterile H2O, on ice.
Electrocompetent bacteria, frozen (50 - 100 µL aliquots), on ice.
BioRad cuvettes (1 mm gap), on ice.
Sterile L-Broth (L.B.)
Bacteria culture tubes
P1000, P200, P20 Pipetman and sterile tips
Sterile Pasteur pipets and bulb.
37ºC incubator and rotating wheel
L.B./Amp bacterial culture plates (or other suitable selection media)
X-gal 20 mg/mL in DMF (dimethlyformamide), store at -20ºC
IPTG 0.2 g/mL in H2O, store at -20ºC

P. Olivier — 1/2000

Solutions

Vortexing pellets while adding solutions is important.

Fixation of Cells

1. Trypsinize cells. Resuspend in PBS containing 5% serum (filtered to remove serum precipitate). Break up clumps by pipetting and spin down cells in 15 mL conical tubes.
2. Aspirate supe and vortex pellet. While vortexing, add 1.5 mL of cold PBS. While vortexing, slowly add 3 mL of cold 95% ethanol in a steady stream. Continue to vortex until thoroughly mixed. At this point samples may be stored at 4ºC for several days or longer.

Staining Procedure

Spins are done in a clinical tabletop centrifuge 5' at top speed, approx 1000 X g

1. Spin down cells, aspirate supe and vortex pellet.
2. While vortexing, add 3 mL of 0.08% pepsin. Incubate at 37°C for 20 min with occasional mixing.
3. Spin down nuclei (expect a very small pellet), aspirate supe and vortex pellet. While vortexing add 1.5 mL of 2M HCl. Incubate 20 min at 37°C with occasional mixing.
4. While vortexing add 3 mL 0.1M Na-borate. Spin down nuclei.
5. Aspirate supe and vortex pellet. While vortexing add 2 mL IFA/Tween 20.
6. Spin down nuclei, aspirate supernatant and vortex pellet. Add 75 µL of 1:5 dilution in IFA of anti-BrdU-FITC and incubate on ice in the dark for 30 min.
7. Add 2 mL IFA/Tween 20 while vortexing. Spin down nuclei, aspirate supe and vortex pellet. Resuspend in 0.25 mL IFA.
8. Add 0.25 mL of 100 µg/mL propidium iodide (in PBS) and incubate on ice in the dark for 60 min. Samples can be stored overnight (even a day or so is OK).
9. Run on FACScan.

How to perform automated counts of fluorescently stained cells.
M. Fero (7/2004)

Note
This protocol describes semi-automated cell counts using fluorescently labeled cells, a hemocytometer and ImageJ software. The hemocytometer is not needed if you have already calibrated the areas of images taken with your microscope and camera. Our setup automatically imports images in iPhoto and loads them into Photoshop with a double-click. However, you could also import and crop the images directly into ImageJ if you prefer.

Materials

• Cells in media
• EtOH
• 1x P.I. (100 µg propidium iodide / mL PBS)
• Hemocytometer
• U.V. fluorescent microscope with camera
• Computer with ImageJ and image cropping software.

Procedure

Fix Cells by diluting them 50% in EtOH.

1. To an eppendorf add 0.1 mL cells in PBS or media
2. Add 0.1 mL of 100% EtOH while vortexing.
3. Spin at 3,000 RPM x1 min.
4. Aspirate off supernatant and respend cells in 0.1 mL of 1x P.I. Mix, incubate 2 min.

Photograph cells on u.v. microscope

5. Add 10 µL of stained cells to Hemocytometer.
6. Place on U.V. microscope. (If the cells are too crowded then you may need to further dilute them in P.I. or PBS)
7. Photograph 1 large square of hemocytometer with brightfield.
8. Without moving the stage or changing the zoom, photograph the same area under u.v. light with red filter.

Import and crop photo (e.g. using iPhoto and Photoshop)

9. Import photos from the camera to the computer (e.g. with iPhoto). Double click an image to open image in Photoshop.
10. In Photoshop, use the marquee tool measure the size of large square of the brightfield hemocytometer image.
    (A large square on the hemocytometer is 1mm x 1mm x 0.1mm = 0.1 µL)
11. Crop the u.v. fluorescent image of cells to same size as a large square.
12. Convert to grayscale (Image > Mode > Grayscale).
13. Save file as .jpg image (medium resolution).

Count cells in cropped image using ImageJ

14. Open .jpg file with ImageJ software.
15. Convert to binary image (Process > Binary > Threshold)
16. Count cells (Analyze > Analyzed Particles... check Display results, Clear results table, Summarize).
17. Repeat counts with additional large squares to be sure that the cells are evenly distributed and to minimize stochastic error. (The standard deviation is ~ sqrt(mean)).

Note: An alternative to cropping the image is to instead photograph the fluorescent cells with a low level of brightfield illuminescense in order to visualize the lines of the hemocytometer together with the fluorescent cells. Select the area of the large square for cell counts directly in ImageJ.

Calculate the Cell Density

Cell density (cells/mL) = Average Cell Count (cells/large square) * 10,000 (large squares/mL) * d.f. (dilution factor, if any)

S. Coats 1995

Materials

P.I. Solution:
4 mM Na3Citrate (0.118 g/100 mL)
30 U/mL RNAseI (43 mg/100 mL)
0.1% Triton-X100 (0.1mL/100 mL)
50 µg/mL propidium iodide (5 mg/100 mL)

Procedure

1. Harvest cells: Rinse with a subconfluent 10 mL dish with PBS (Ca++/Mg++ free).  Cover with 1.5 mL 0.1% (2.5 mM) EDTA in PBS at 37ºC x 10 min.  Loosen cells by vigorous pipetting then transfer suspension to 1.5 mL Eppendorf tubes on ice.
2. Spin at 1000 RPM, 4ºC.  Discard supernatant.  Resuspend in 0.5 mL PBS.  Remove 0.1 mL for flow cytometry and use the remainder for protein extracts.
3. Add 4 - 5 volumes of 100% EtOH and vortex gently.  Incubate 15'. (The cells may be now stored at 4ºC.)
4. Spin at 1000 RPM, and wash pellet with PBS.
5. Resuspend pellet in 0.5 mL of P.I. solution (above).  This should give ~1 million cells/mL.  Incubate 10 min. at 37ºC.
6. Filter through nylon mesh screen to remove cell clumps.  Keep at 4ºC on the dark until ready for flow. See the Flow Cytometer Protocol for more details.

M. Fero — 3/02

Protocol

1. Wash cells with PBS and trypsinize to a single cell suspension.
2. Count an aliquot on a hemocytometer. Meanwhile centrifuge the cells (e.g. 1000 RPM, 5 min.) to pellet them.
3. Resuspend the cells to a concentration of 2 x 106 cells/sample with Flow Sort solution.
4. Aliquot into flow sort tubes (maximum 3 mL per tube)
5. Aliquot 1.5 mL of media into falcon tubes for collection.
6. Place cells and collection tubes on ice and transport to flow facility. Also bring transfer pipets and extra tubes.
7. Run untransfected cells first and set up voltage for GFP (FL1) and PI for viability (FL3).
8. Sort GFP(+), PI (-) cells. The maximum sort rate is about 5,000 total cells/second.
9. Sort for a maximum of one hour.
10. Spin down cells and plate (104/cm2 or 106 cells/P100).

Materials

• GFP transfected cells
• Untransfected cells as negative control
• Flow Sort solution: 25% media, 75% PBS with 2µg/mL propidium iodide, 10 µg/mL DNAse (or 10 U/mL)
• PBS
• 1x Trypsin/EDTA
• Media
• Flow sort tubes: Falcon #352054
• Sterile transfer pipets

J. Orthel.  6/27/03
mf revised 9/04

Materials

Capillary tubes
1.5 mL Eppendorf microfuge tubes
15 mL conical centrifuge tubes
96-well V-bottom plates (Corning Costar 3894, from Fisher)
Flow tubes (Falcon 352054)

Reagents

0.5 M EDTA pH 8.0
PBS + 0.5% heat inactivated FBS

RBC Lysis Buffer
4.15 g NH4Cl
0.5 g NaHCO3
0.0186 g Disodium EDTA
200 mL H2O

Procedure

1. Collect 200 - 500 uL blood from each mouse.  Mix with 50 uL 0.5M EDTA in 1.5 mL eppendorf tubes.
2. Add 250 uL blood to 5 mL RBC Lysis Buffer (20x vol. blood) in 15 mL conical tubes.
3. Spin down white cells 1500 RPM x2 min.
4. Aspirate lysate and wash by resuspending cells in 5 mL PBS/FBS.
5. Spin down at 1500 RPM x2 min.
6. Resuspending cells in 500 uL PBS/FBS with 1 uL Fc Block (1/500)
7. Add 150 uL to each well of a 96 well V-bottom dish.
8. Add 50 uL 1º Ab Master Mix (the mix is a 1/25 dilution of each 1º Ab in PBS/FBS).
9. Include 1 well with a combination of Isotype controls for setting voltage.  Also include 1 well for each of the 1º Ab as single positive controls for setting compensation.
10. Incubate 60 min at 4ºC.
11. Spin 1500 RPM x2 min.  Discard supernatant by shaking it out once into the sink, and blot inverted plate on paper towel.
12. Wash by adding 200 uL PBS/FBS to each well, and mix by pipetting up and down.
13. Immediately spin at 1500 RPM x2 min. and discard supernatant.
14. Add 150 uL of 2º Ab (i.e. Streptavidin-TC) Master Mix.
15. Incubate 30 min at 4ºC, then spin at 1500 RPM x5 min.  Shake out supernatant.
16. Resuspend in 200 - 500 uL of PBS/FBS and transfer to 5 mL flow tubes.
17. See the flow cytometry protocol for 3-color flow.

Dan Kuppers 5/2011
(ref. Rothaeusler, K. and Baumgarth, N., Cytometry Part A 2006)

Materials

• BrdU 10 mg/mL, store frozen
• 10 million in vivo labeled thymocytes
• PBS w/o Ca or Mg
• PBS w/ Ca/Mg (PBS with 0.1 g/L CaCl2, 0.1 g/L MgCl2•6H2O)
• Wash Solution: PBS + 0.5% NP40
• Fixative: Fresh 1% paraformaldehyde + 0.05% NP40
• Quick 1% paraformaldehyde:

1. Mix 0.1g paraformaldehyde w/ 0.5mL H2O and one drop of 1N NaOH
2. Heat 2-3 min @ 80ºC untill disolved.
3. Add to 9.5 mL 1x PBS
4. Check that the pH is 7.4 (should not require adjusting)

• Cell surface antibody (e.g. PE conjugated Pharmingen antibodies diluted 1:100 - 1:50 in PBS)
• DNAse I (Invitrogen 212 U/µL)
• FITC-conjugated anti-BrdU antibody (Sigma, clone BU-33)
• Cell surface antibodies of interest
• 15 mL conical tubes
• Flow Cytometry tubes

Procedure

1. Label cells in vivo by injecting mice with 1mg of BrdU, i.p. 1 hr. prior to harvest. Include one no-BrdU mouse as a negative control.
2. Prepare a single cell suspension in PBS and count cells on a hemocytometer. Aliquot out 10 million thymocytes and spin at 1000-1500 rpm x5 min to pellet.
3. Discard supernatant and resuspend cells in 200 µL solution of conjugated cell surface antibodies of interest (1:50 - 1:100 in PBS) @ 4°C for 1hr. Note: Novel fluorophores should be be tested for loss of signal from fixation.
4. Wash 2x with PBS. If biotinylated antibodies were used then incubate cells in 200 µL of streptavidin-APC (1:100 in PBS).
5. Wash cells 2x with PBS. Loosen pellet and then incubate cells in 1mL Fixative o/n @ 4°C.

Next day

1. Wash cells 2x with PBS.
2. Resuspend cells in 250 µL PBS + Ca/Mg with 50 U/mL DNase I. Incubate @ 37°C for 30 min.
3. Wash 2X with Wash Solution and resuspend in 100 µL of a 1:5 PBS solution of FITC-conjugated anti-BrdU antibody. Incubate on ice for 45 min.
4. Wash with 1mL Wash Solution and resuspend the cells in 0.2 mL PBS (optionally with 10 µg/mL DAPI or PI) and transfer to flow cytometry tubes. If DAPI or PI is used incubate @ 4°C for 30 min. prior to flow cytometry.
5. Analyze the cells by Flow Cytometry. Use the no-BrdU control sample to set voltages. This is important since the anti-BrdU antibody gives a high background.

M. Fero — 6/01 and D.B. — 12/98

Protocol

1. Harvest 106 cells/sample with PBS/FBS into a 15 mL conical tube spin at 1300 RPM. Wash by resuspending cells in 1.5 mL of PBS/FBS by spinning at 3000 RPM in a microcentrifuge.
2. Incubate with 100 µL (use Pharmacia antibodies at 1:500) of primary cell surface antibody in PBS/FBS for 30 min. (Keep in dark if it is labeled). Wash with PBS/FBS.
3. If the primary antibody is unlabled incubate with FITC or PE-conjugated secondary antibody if necessary for 30'. Wash in PBS/FBS.
4. Fix cells with 1 mL of 0.5% Paraformaldehyde in PBS for 5 min. r.t and mix several times. Wash with PBS/FBS.
5. Fix cells in 1.5 mL of 50% EtOH. Incubate 30 min. Spin.
6. Wash in 1 mL of 1 mL of Hypotonic solution, spin. Wash with PBS, spin.
7. Incubate in 100 µL of primary (intracellular) antibody in PBS/FBS for >30 min. Wash with PBS/FBS.
8. For an unlabled rabbit primary antibody add 0.5 mL biotin-conjugated anti-rabbit secondary antibody (1:10,000) for 30 min. Wash in PBS/FBS.
9. For biotinylated primary antibodies incubate in 100 µL Steptavidin-TriColor (1:100 in PBS/FBS), 30 min. Wash with PBS/FBS.
10. For DNA content add 500 µL of 0.5 µg/mL Höchst 33342 (LSR, FL-5), or 0.5 µg/mL DAPI (LSR, FL-5), or PI (propidium iodide, 10 µg/mL)

Materials

• PBS/FBS: 0.5% heat inactivated (56°C, 30') FBS in PBS.
• 0.5% Paraformaldehyde in PBS
• 50% EtOH
• Hypotonic Solution: 10 mM HEPES, 0.55% NaCl, 0.1% NaN3, 4% h.i.FBS.
• FITC (FL-1) or PE (FL-2) conjugated cell surface primary antibody
• Primary nuclear antibody
• Biotinylated secondary (Goat anti-rabbit, Caltag L42015)
• Streptavidin-TriColor (Caltag, SA1006) (FL-3)
• Höchst 33342 (or DAPI) 0.5 µg/mL for DNA staining.

M. Fero — 12/02

Reservations

Schedule time on a cytometer (FACS or Calibur) by filling in the calendar at the flow lab. Don't grossly overbook since we have to pay for both reserved time and the time actually used (which ever is greater).

Prepare Cells

Cells should be stained ahead of time and suspended as single cells in PBS to a density of about 20,000 to 100,000 cells/mL in flow tubes (see separate staining protocols). Pass cells through nylon mesh if they may have clumps (not necessary for hematopoietic cells). Bring to the flow lab: Cells in flow tubes, extra PBS, listing of samples. A negative control should always be included for setting up the machine voltages. The ideal negative control for antibody staining is a no antigen control (e.g. no BrdU added to the culture dish, or a knockout cell line). If this is not available then cells stainined with pre-immune serum + secondary or flourescently labeled isotype antibodies should be used. If multicolor flow is being used multiple isotype antibodies may be used on a single negative control specimen. For multi-color flow, positive controls stained with a single antibody should also be employed to allow the setting of compensation (see below).

Start Machines

Fill the sheath fluid reservoir of the cytometer with PBS before getting started. The Beckman Calibur cytometers have a green power button on the right side. There is also a second power toggle switch hidden on the back of the machine. Boot the computer after the cytometer is turned on (otherwise the BD startup init won't "find" the cytometer).

Launch the CELLQuest application (an alias is under the apple menu items).

Open your template. If you wish to use a previous template close the blank window and open the template which should be stored in the templates folder. (Desktop > Templates > Fero > ... ) It is also a good idea to save a copy of your templates on one of our computers so you can transfer it to a different cytometer in the future if necessary. The template is simply a program which lays out your acquisition windows and gate parameters. It does not contain any data nor does it affects the cytometers voltage and compensation settings.

Select "Connect to Cytometer" from the Acquire menu. If this is dimmed then the program does not realize that you are attached to a cytometer. Try relaunching the program or if necessary reboot the computer.

Select "Intstrument Settings" from the Cytometer menu. (This option doesn't appear until you are connected to the cytometer). If you wish to use previous instrument settings then select "Open" in the Insrument Settings dialogue box. Navigate to the settings file you wish to use. (Desktop > Instrument Settings > Fero > ) Press the "Set" button to make the settings take effect and then "Done" to close the dialogue box. Note: Instrument settings can not be opened directly from the Finder where they normally will have a "blank" appearing icon. If you change the voltage or compensation settings during your flow session you may want to save these settings with a new name for future use.

Select "Acquisition and Storage" from the Acquire menu. The following settings are recommended: Acquire = All, Event count = 10,000, Storage = All, Resolution = 1024, Parameters saved = (check that the appropriate Photomultiplier tubes are checked). Close window.

Select "Parameter Description" from the Acquire menu. Select the "Folder" button and navigate to the place you will store your new data files (Desktop > Data Files > Day of Week) Select the "File" button. Enter a prefix in the following format MMDDYYc2 (where c2 is for calibur 2 and c1 is for calibur 1). Look at the notebook next to the cytometer to see which file number you should start with and enter this value. Close the File name window but keep the Parameter Description window open (but you can cover up the bottom half). It is a good idea to enter the sample ID values for each sample and to also record the file numbers in your notes.

Open "Detector/Amps" , "Compensation" and "Threshold" from the Cytometer menu. Be sure the threshold is set to an FSC of 50 and then close this window. Open the "Counter" window under the Acquire menu.

Creating Templates

If you aren't using a premade template then you should use the tool palette or the "Plots" menu to create FSC vs SSC Dot Plots and Histogram plots of FL1, FL2, or FL3 as necessary. If multicolor flow is being performed then Dot plots of FL1 vs FL2, etc. should also be set up to aid in adjusting the compensation (see below). If a data plot is already created, it can be changed by selecting it and then typing Command+F (or select "Format Histogram.." from the Data menu.) Usually gates should not be used while initially setting up the voltage or compensation.

Voltage Adjustment

On the Acquisition Window check the "Setup" button so you're not saving data. Vortex your negative control specimen. (Ideally this is a sample of control cells which has been stained with a mix of fluorescently tagged isotype antibodies corresponding to the antibodies you will be using in your analysis).

FSC and SSC: While viewing a dot plot of FSC vs SSC adjust the voltage of FSC and SSC photomultiplier tubes until the bulk of cells appear roughly in the lower left quadrant.

FL1: Viewing a histogram of FL1 (with no gate) adjust the FL1 voltage. Be sure that "Log" not "Lin" is selected in the Voltage window. Adjust the voltage until you get a peak in the FL1 histogram between 100 and 101

This process should be repeated for the remaining photomultiplier tubes:

FL1: FITC, GFP, YFP, Hoechst
FL2: Phycoerythrein (PE), Propidium Iodide (DNA content)
FL3: TriColor, Propidium Iodide (viability)
FL4: DAPI (LSR machine)

Note if you are using P.I. for a DNA content analysis then FL2 should be set to display on "Lin" instead of "Log".

Compensation Adjustment

If you are doing flow with multiple colors you should also adjust the compensation values to minimize the "spill over" of one color into another channel. Place a FITC-only positive control sample on the machine. Using a dot plot of FL1 vs FL2 be sure that cells positive for FL1 do not exceed a value of 101 for FL2. Use the minimum amount of compensation of FL2 (FL2 = FL2 - %FL1) to eliminate any spill over of FL1 into the FL2 channel. Note: Decreasing the voltage on FL1 may also be necessary to prevent spillover into FL2 if the flourochrome is too bright or has a broad spectrum.

If the FL2 channel is being used this process should be repeated with a different single positive control (e.g. PE labeled antibody). Use a dot plot of FL2 vs FL1 to adjust the spill over of FL2 into FL1 (using the FL1 = FL1 - %FL2 compensation slider). A second dot plot displaying FL2 vs FL3 should be used to adjust the spill over of FL2 into FL3 (using the FL3 = FL3 - %FL2 compensation slider).

If the FL3 channel is being used (e.g. TriColor) then one should compensate for the spillover of FL3 into FL2. Use the dot plot of FL2 and FL3 for this with a single positive control stained with the TriColor antibody. Similarly adjust the compensation slider (FL2 = FL2 - %FL3) until the signal in FL2 < 101.

Gating
Gating will allow you to view cells of interest by any combination of criteria that you choose. Gating does not change the intensity value assigned to an event as is the case for changes in voltage or compensation. It simply lets you decide which data to view and which data to ignore or discard. It is important to check that small changes in your gates don't have significant effects on your results or else your data will be prone to artifact. When you create or format a data plot (i.e. on the Data menu select "Format Histogram" or "Format Dot Plot") you can select any of the gates that you have created. This will filter the data and plot will only display those events which meet the gate criteria. Gating will not discard data unless you have requested this under "Acquisiton and Storage". Gating can subsequently be changed when you analyze your data without any loss of information.

To set up a gate you first draw a "Region" using one of the tools on the tool palette (there are four geometric shapes to choose from outlined with dotted lines). Note that the "Marker bar" (designated with an M) and the quadrant maker tool next do not define regions. They are used for statistical analysis only and can't be used to filter data like the regions/gates.

Regions which are commonly employed include:
PI for DNA content: FL2 area vs FL2 width. This window is useful for gaiting out apoptotic cells (lower left quadrant) and doublets (a separate cloud with increased FL2 width).
FSC vs SSC: This is useful for gating out RBC, myeloid cells etc, from blood or marrow.

Each region that is setup automatically defines a gate (e.g. G1 = R1). To delete regions select "Region List" under the Gates menu. To combine regions into more complex gating criteria use the "Gate List" under the Gates menu. (e.g. G5 = (R1 or R2) and R3).

Acquiring Data

In the "Acquisition" window deselect the "Setup" button. Now when you select "Acquire" the cytometer will collect 10,000 events and automatically save a data file with the location and naming convention that you previously specified. Vortex each sample before running. Select a flow rate button on the machine (Low, Med, or High) to keep the event rate between 200 ­ 1000 events per second. The maximum event rates are around 4000/sec, attempts to increase the flow rates in this case will actually reduce the event rate by increasing the percentage of events that the cytometer rejects. Enter the Sample ID values if you wish these to be stored in each data file. (These can later be viewed under "Histogram Stats" for each data file).

When you finish running your samples consider saving your template for future analysis and consider saving your instrument settings. Also consider moving your data files and template to the lab computer for analysis. Print out your histograms for your notebook.

Troubleshooting
If you get a "cytometer not ready" you probably left the cytometer on "standby". Otherwise you might be running out of sheath fluid. Check the reservoir directly or select "Status" under the Cytometer menu. Don't let the sheath fluid run dry or you'll have trouble from air bubbles clogging the cytometer.

Shut Down

The cytometer should be left on in "Standby" mode with a tube of water on the intake nozzle when not in use. If you are the last person scheduled to use the cytometer you should follow the shutdown procedures listed in the flow lab. Basically this consists of running dilute bleach, detergent and then water through the machine each for 5 minutes and then power down both the cytometer and the computer.

Sign Out
Record the file numbers that you used in the log book by the cytometer so the next user doesn't overwrite your data files by accident. Also sign out at the computer indicating the budget number and the time that you used the machine. It's important to do this correctly for your data files to be transferred to your Fred account.

Analyzing Data

Stored data files may be analyzed after acquisition is complete. You may use your previous template to make matters easier but this is not necessary. From an open template select an individual histogram or data plot and select "Format Histogram (Command+F)". From the dialogue box you must change the selection from Acquisition to Analysis. Select the data file you wish to open. Repeat this process for each histogram or dataplot in the template. To increment the histograms to the next data file select them and then press (Command + ] ) or "Next data file". Statistics may also be viewed on selected histograms, and histograms may be overlayed on on another in the analysis mode.

(M. Fero 7.10.06)

Materials

• Trypsin (Gibco 25200-023)
• 3T3 Medium: 500 mL DME (Invitrogen) + 50 mL FBS (Hyclone) + 5 mL 100x Pen/Strep
• 2x Freezing Medium: 3T3 Medium + 20% DMSO: Filter sterilized.
• Sterile PBS
• P100 and P60 tissue culture dishes
• Dissecting tools: Scissors, forceps, scalpel
• 95% ethanol in a squeeze bottle and in a beaker
• Mouse breeders

Procedure

Note: This protocol describes the isolation of primary MEFs and passage according to the 3T3 protocol. Passaging primary cells on the 3T3 protocol will maximize the growth prior to the development of cellular senescence. The 3T3 protocol was originally described as the passage of 3 x 105 cells every 3 days on 50 mm. dishes which is equivalent to 1.2 x 106 cells on a P100 dish (Nilausen K, Green H. Exp Cell Res. 1965). If cells are passaged continuously every three days under normoxic conditions they will experience a period of rapid growth (passage 1 - 5), progressively slower growth (passage 5 - 10) and senescence (little or no growth, passage 10 - 25). After this time the a subset of cells will emerge from crisis, through the selection for a more rapidly growing, immortalized subclone. This is typically accompanied by the development of an aneuploid karyotype. Early passage primary MEFs and immortalized MEFs are usually spindle shaped and form densely packed monolayers, whereas senescent cells tend to flatten and spread out. Some lines may become transformed and display neuron-like or refractile morphology, growth despite serum starvation, growth in suspension or in soft agar, and the ability to form fibrosarcoma tumors in mice. Senesence in murine fibroblasts is a result of oxidative stress as growth in physiologic (3%) O2 allows continual growth of murine cells similar to human fibroblasts (Parrinello S., et al. Nature Cell Biol, 2003).

A. Isolating Primary MEFs:

1. Setup mouse breeder pairs in the afternoon. Ideally one male should be setup with one or two virgin 7-8 week old females.
2. Check females for copulatory plugs the following morning. If present this should be marked as day +0.5 (or round off to day +1). Remove plugged females from the males breeder cages. Check the remaining females on a daily basis.
3. Embryos should be harvested on day 11.5 to 13.5 as follows:
   1. Place clean forceps and scissors into a beaker with ethanol. Fill a second beaker with sterile water or PBS to wash instruments between embryos. Aliquot sterile PBS into P60 dishes (2 - 3 per embryo).
   2. Euthanize the pregnant female according to standard protocols. CO2 inhalation should be avoided because this induces acidosis.
   3. Saturate fur with ethanol. Incise the skin vertically and pin to the side.
   4. Using a fresh pair of forceps and scissors open the body wall and pin this back as well.
   5. Extract the intact uterus by grasping one horn with forceps. Withdraw the uterine horn from the abdomen without letting it touching skin or non-sterile areas. Cut the cervix and blood vessels and and transfer the uterus to a sterile petri dish.
   6. The embryos will appear likes pearls on a string. Cut the uterus into sections between each embyo. Place a uterine section with a single embryo into a fresh petri dish with sterile PBS. Squeeze the uterus with forceps to expell the embryo through the cut portion of the uterus.
   7. Wash the embryo in a fresh dish with sterile PBS to wash away any remaining maternal blood cells.
   8. Transfer the embryo to a fresh Petri dish with sterile PBS. Remove the membranes and umbilical cord.
   9. With a scalpel vertically incise the abdominal wall and remove the pink hematopoietic tissue (liver and spleen) as well as the tubular intestine.
   10. Trim off the bulk of the CNS tissue by dissecing away the head above the level of the oral cavity.
4. Save CNS or liver tissue in a 1.5 mL tube for DNA extraction and genotyping. Embryos should be signed a mouse ID and genotyped according to standard protocols. It may be necessary to dilute the DNA or limit the PCR cycle number because of the large amount of DNA and the possibility of maternal blood contamination. Transfer the embryo body to the tissue culture hood in a P100 dish with 5 mL of 1x trypsin.
5. Mince each embryo for 2 - 3 min. with a sterile forceps and scissors or a scalpel to create bits smaller than 2 mm. Place the dish in a 37ºC incubator. Rinse the instruments in sterile water and then 95% ethanol to clean and air dry them between each embryo.
6. After a total of 15 minutes transfer the cells and tissue bits with a 10 mL pipet to a 15 mL conical tube. Pipet the cells several times to further dissociate tissue bits. Add 5 mL of media (DME + 10% FBS).
7. Spin at 1000 RPM in a table top centrifuge for 5 min.
8. Aspirate the supernatant. Resuspend the embryonic cell pellet in 10 mL fresh media and transfer to a clean sterile P100 dish. Small bits of tissue in the cell suspension are OK. MEFs will migrate out of tissue bits onto the dish.
9. Mark the plates as "passage 0", with the date, and the embryo ID. Evenly distribute the cells and tissue bits by horizontal agitation and then incubate at 37ºC in 5% CO2. (Optional: To minimize senescence grow cells in 3% O2, see above).

B. Passaging primary MEFS:

1. Check the plates daily until they are nearly confluent (usually 1-3 days).
2. To trypsinize the cells:
   1. Aspirate away the media. Add 5 mL of PBS. Swirl to rinse and then aspirate off the PBS.
   2. Add 3-4 mL of 1x trypsin (or enough to cover the plate). Incubate at 37ºC for 2 min.
   3. Tap the plate to dislodge cells. Continue incubating until start to come off in large sheets. (For the 1st passage it is sufficient to loosen up the cells in the monolayer. Do not dislodge larger chunks of tissue, if present.)
   4. Add 5 mL of media (DME + 10% FBS) and transfer to a 15 mL conical tube. Pipet to create a single cell suspension. (If larger chunks of tissue are present in the first passage - allow them to settle out of suspension for 1 minute. Transfer the suspended cells to a fresh conical tube.)
3. Determine the viable cell count:
   1. Add 10 µL of cells suspension to a multiwell dish. Add 10 µL of 2x Trypan Blue stain.
   2. Mix by pipetting and transfer 10 µL to each side of a hemocytometer.
   3. Count white (live) and blue (dead) cells on each side of the hemocytometer. Count a total of 100 - 200 cells to be reasonably accurate.
   4. Multiply the average number of live cells in each large square x104 to calculate the number of cells per mL.
   5. Keep a record the number of cells present and amount that they will be diluted to be able to calculate doubling times.
4. Meanwhile spin the remaining cells for 5 min. at 1000 RPM.
5. Aspirate off the supernatant leaving ~ 0.2 mL of media behind. Resuspend pellet by flicking bottom of tube. Add fresh media to bring the cells to a density of 107 cells / mL.
6. Add 9 mL of media to each 100mm dish + 1 mL of cell suspension to plate 1.2 x 106 cells per dish.
7. Mix the cells on the dish by swirling the plates. Horizontally agitate the plates in the incubator to ensure an even distribution of cells. Incubate 37ºC + 5% CO2. (Optional: To minimize senescence grow cells in 3% O2.)
8. Passage the cells every 3 days as described above (B.2-B.7) even if there is little or no growth since the last passage.

C. Freezing/thawing MEFS:

1. Primary MEFs do not do freezer and thaw well and are likely senesce at a lower passage than are cells kept in continuous culture. If they are to be frozen it should probably be at passage 1 - 2.
2. To freeze MEFs:
   1. Trypsinize cells as above B.2 - B.4. Resuspend the cells at a density of 4 x 106 live cells /mL. Add an equal volume of 2x Freezing Media (media + 20% DMSO).
   2. Aliquot 1 mL of cell suspension in freezing media into cryo vials. Cap securely, being careful not to contaminate the edges of the vial or cap. Label tubes as MEFs, passage #, date, and your initials.
   3. Place vials in a styrofoam container or a cell freezing jar with isopropanol. Place at -80ºC overnight.
   4. Transfer vials to a cryofreezer. Record the location in the Freezer database.
3. To thaw MEFs:
   1. Pre-warm medium at 37ºC.
   2. Remove a vial of cells from the cryo freezer and place in 37ºC bath.
   3. Delete the vial entry from the Freezer database.
   4. Wipe the vial with ethanol and place in the tissue culture hood.
   5. Transfer the cells to a sterile conical tube with 10 mL of medium.
   6. Count the viable cells with trypan blue as above (B.3).
   7. Plate the cells at density of 1.2 x 106 cells per P100 (or 1.5 x 104 cells / cm2 in a smaller dish).
   8. Incubate 37ºC + 5% CO2. (Optional: To minimize senescence grow cells in 3% O2.)

(M. Fero — 1/4/95, with thanks to P. Soriano)

MATERIALS

Glassware: All hand washed with no soap.

Gelatin (Sigma G-1393) Diluted to 0.1% in H2O, + 1 drop of media to colorize

Plastic pipets, P100 tissue culture dishes and 24 well multiwell dishes.

Trypsin (Gibco 25200-023)

BioRad cuvettes (0.4 cm, electrode gap 50, Cat# 165.2088)

PBS (Dilute from a 10 Stock solution, autoclaved)

DME:

1 pk Gibco DME powder (cat #: 12100-046)

2.2 gm NaHCO3

q.s. to 1 L with millipore H2O

pH to 7.3 with ~ 5 drops of 12M HCl

7.5 µL ß-ME

5 mL Pen/Strep (Stock = 10K u/mL Pen + 10 mg/mL Strep)

+ 10 mL 200 mM Glutamine (if older than 1 mos.)

STO cell media: DME + 10% FBS

ES cell media: DME + 15% FBS (lot titered for ES cell toxicity)

2x Freezing media:

30 mL DME

10 mL FBS

10 mL sterile DMSO (Sigma D 2650), final concentration of 20%.

Cryovials and cryo-freezing container

MMC, mitomycin C (Sigma M-0503) 0.5 mg/mL in PBS, filter sterilized, light protected, 4°C

Glass tubing (Kimble 1.2 - 1.5 mm borosilicate glass)

Cell lysis buffer:

KG buffer with no BSA or gelatin

1% ß-ME instead of 0.5% ß-ME

0.5% Triton X-100

proteinase K 0.5 mg/mL.

LIF and Neo (or Hygro) transformed STO cells

E.S. cells

PROCEDURE

Splitting STO cells:

1. Aspirate off media.
2. Rinse with 6 mL of PBS, and aspirate off.
3. Add 1 mL of trypsin, 37°C x 3 min.
4. Add 2 mL of media to plate, and loosen up cells with a repeated pipeting. Save 1 drop to quantitate in hemocytometer while spinning.
5. Spin 5 min. Suck off supernatant and resuspend cell pellet in minimal volume by flicking. Add 1-2 mL of media.
6. Plate 5 x 10^4 cells/cm2 (8.8 x 10^6 cells) onto 5 large 150 mm plates with 20 mL of media. (10^-4 mL/large square in hemocytometer)

Freezing STO cells:

1. Trypsinize cells as when splitting them.
2. Resuspend to a concentration of 4 million cells per mL in STO cell media.
3. Add 1 volume of freezing media.
4. Aliquot 1 mL into cryo vials.
5. Gradually cool to -20°C in a cryo container for 2 to 4 hrs.
6. Transfer to liquid N2 for storage.

Mitomicin C inactivation of STO cells:

1. Gelatinize and appropriate number of 100 mm plates (approximately 60 plates total will be needed to target a single ES cell construct.) Add 5 mL of gelatin solution to each plate for 20 min. Aspirate off the gelatin and allow to air dry.
2. Add 0.4 mL of MMC (0.5 mg/mL stock) in 20 mL of fresh STO media to each 150 mm plate of subconfluent STO cells
3. Incubate at 37°C for 2 to 4 hours.
4. Rinse off MMC with PBS twice. Trypsinize and plate STO cells onto the gelatinized plates in 8 mL of media at the same density listed for splitting cells, above (4 million/100 mm plate).
5. Label plates with the day of the month.
6. Feed cells every 2 weeks. They are good for 1 month.

Thawing E.S. cells:

1. Gelatinize a 100 mm and two 150 mm plates by covering with gelatin solution for > 1 hr. (Keep 150 mm plates for passage of STO cells.
2. Quickly thaw vial of STO cells at 37°C.
3. Gradualy add thawed cells to media in a 15 mL conical tube. (While gently agitating). Spin cells to pellet, then resuspend in 1 mL of media. Count with hemocytometer. (10^-4 mL/Large square) Use 5 x 10⁴ cells/cm2, or 4 x 10⁶ cells per 100 mm dish.
4. Add cells to 100 mm dish with 8 mL of STO media.
5. Incubate 37°C, 5% CO2, o.n.
6. Split cells the following day.

Splitting E.S. cells:

1. Warm media and thaw trypsin.
2. Aspirate off media. Rinse cells 1x with PBS.
3. Cover cells with 0.5 mL trypsin. Incubate at 37°C for 5 min.
4. Agitate plate and incubate at 37°C for 1 additional min.
5. Break up cell clumps by repeated pipetting and add 0.5 mL of E.S. media to stop enzyme.
6. Count cells on hemocytometer (10^-4 mL / large square) and pellet remainder of cells at 1000 rpm.
7. Plate 5 million cells/P100 dish (75 cm²) in 10 mL of E.S. media.

Electroportating E.S. cells:

1. Linearize 100 µg of the DNA construct and inactivate the restriction enzyme. Ethanol precipitate the DNA and resuspend it at a concentration to 2.5 mg/mL according to a flourimeter. (This probably corresponds to 5 mg/mL on a spectrophotometer).
2. Refeed ES cells in a.m.
3. Rinse off media with PBS.
4. Add 2.5 mL of trypsin, incubate at 37°C for 5 min. Agitate dish and incubate for 1 min. more.
5. Inactivate trypsin with 1 mL of media. Resuspend well with a transfer pipette.
6. Count cells with a hemocytometer and pellet cells at 1K rpm.
7. Resuspend cells at 12.5 x10⁶ cells/ mL in PBS. (Note: Previously listed as 1.25)
8. Transfer 0.8 mL to a BioRad cuvette. Add 10 µL of (2.5 mg/mL) DNA. Electroporate at 230 V. 500 µFD which should give a time constant of 5 msec.
9. Transfer cells to 10 mL of ES media and plate on a single P100 plate of STO cells (for promorless constructs) or on 6 plates (for constructs containing PGK-Neo).
10. On the following day, and daily therafter feed the cells with ES media supplemented with G418 (300 µg/mL) for Neo constructs, or hygromycin at 150 µg/mL. Also supplement with gancyclovir (2.5 µM, MW=255) if an HSV-TK construct is being utilized. Treat one plate with G418 alone to test the effectiveness of the TK selection.  Feed cells daily with fresh pre-warmed media with the appropriate antibiotics for at least one week or until colonies clearly emerge (~10 days).

Note:  If you are using a fresh batch of antibiotics or a new selection vector it may help to emperically determine the ideal drug doses.  Conduct a series of 3-fold dilutions bracketing the usual drug concentration for a total of 5 doses (versus no drug).  Electroporate 10⁶ ES cells with an empty selection vector and plate them into a 6 well dish containing inactivated STO cells.  Compare this to an equivalent number of ES cells electroporated with no DNA.  Feed the cells daily, quantify colony formation at 10 days and observe for signs of differentiation of the colonies.  Typically 1/1000 cells will stably but integrate the DNA (or about 160 colonies per well).  If double drug selection is being performed (e.g. Neo + TK genes) the entire titration should be repeated using a constant G418 dose and variable gancyclovir doses using a vector known to target a specific gene.  Ganciclovir typically induces a 10-fold reduction in colony formation with a targeting vector (and an equivalent enrichment of proper integration events).  The efficiency of a targeting vector is exponentially related to the total length of its homology regions.

Picking E.S. cell clones for PCR screening:

1. Prewarm media and PBS.
2. Constuct a mouth pipette (From a mouth piece, rubber tubing, a Drummond 0.8 µm filter, a yellow tip, 1.0 mm I.D. Tygon tubing). Pull glass capillary tubes over a Bunsen burner with a low flame. Break off tubes with a 2" stem.
3. Aspirate the media from plates. Invert the plate on the microscope and dot large clones (2 mm) which are well separated. Number the clones or circle them in groups of 5 to10 if pools are to be screened.
4. Cover the colonies with 10 mL of PBS. Pick off 1/5 of a colony under a dissecting microscope. (Tips: Don't suck up air bubbles, hold the mouth piece in your teeth, puff on the tubing like a cigar to draw up small quantites, open your mouth to break the vacuum and cease drawing up - it will usually stay put due to capillary action. Alternatively, use our tongue to stop the capillary flow).
5. Transfer the colony fragments to eppendorf tubes.
6. After 15 or 20 min of picking recover colonies with warm media and return to the incubator. Alternate plates to let the cells recover.
7. Spin the tubes and remove the PBS. Resuspend cells in 10 µL of Cell Lysis buffer.
8. Alternatively, if single colonies are being picked in a minimal volume they can be transfered directly into tubes containing 12 µL of Cell Lysis buffer. This way there's no need to spin the cells.
9. Lyse the cells by incubating at 55°C for a total of 1 hr. Vortex and spin the tubes to reduce the condensation in the middle and again at the end of the incubation time.
10. Heat denature the DNA at 95°C for 5 min.
11. Use 1 µL of DNA for a 20 µL PCR reaction.

Replating E.S. Clones:

1. Spot out trypsin onto a sterile dish in 7 µL aliquots and cover to minimize evaporation.
2. Aspirate media from the E.S. cells and cover them with 10 mL of PBS.
3. Transfer the colonies of interest using the mouth pipette and a "larger bore" pulled cappillary tube in a minimal volume of PBS to the trypsin droplet.
4. Incubate at 37°C for 5-10 min. The convention is to consider this an additional "passage" when counting passage numbers.
5. Meanwhile give the feeder STO cells fresh E.S. media on 24 well dishes.
6. Using a pipetman and gel loading tips break up the colony with repeated aspirations, and then transfer the cells to the dish of STO cells. Agitate at 90° angles to evenly distribute the cells and incubate at 37°C.
7. Feed cells daily with E.S. media.
8. As the cell number increase transfer to 6 well dishes and then P100s.

M.Fero — May 5, 2003

Materials

4 mice from each genotype
4 Ly5 mice
Buckets with wet ice 3x
Bucket with dry ice 1x
Dewar flask with liquid nitrogen
100 mL beakers with 95% ethanol 2x
100 mL beaker with dd. H2O 2x
Sterile petri dishes
Surgical tools
Sterile IMDM medium aliquoted into 50 mL conical tubes
Sterile Ficoll-Hypaque (d = 1.077)
Halothane in a bell jar
Glass slides
RBC lysis buffer

Procedure

1. Weigh mouse.
2. Anesthetize with halothane or isoflourane inhalation in a bell jar.
3. Place the ends of surgical instruments in a 100 mL beaker with ethanol. Keep a second beaker with sterile water or PBS for washing the instruments. Wet the skin with 70% ethanol. Open the skin and pin back to keep the body wall sterile.
4. Open the chest and aspirate blood from the heart into a 1 mL syringe through a 23G needle. Transfer blood to Eppendorf tube with a drop of EDTA. Mix continually to prevent clotting. Spot 10 µL onto the end of a glass slide and make a thin smear. Stain the smears with HemaStain according to the standard protocol. Perform RBC and WBC counts on hemacytometer. Perform differential counts of 200 WBC on the blood smears.
5. Remove and weigh thymus. Place into 1.5 mL eppendorf tube and plunge into liquid nitrogen. Store on dry ice and then freeze at -80C.
6. Remove spleen and place it in a sterile petri dish, record weight, keep dish on ice.
7. Remove femurs taking care to trim off as much muscle as possible. Keep instruments clean by washing in water or PBS and dipping ends into 95% EtOH.

In cell culture hood:

1. Cut ends off of femurs and flush 1 mL of IMDM through each end into a 15 mL conical tube.
2. Mince spleen with two sterile scalpels in a few mL of IMDM. Transfer suspended cells to 15 mL conical tube.
3. Pool bone marrow from each genotype and load onto Ficoll in two 15 mL conical tubes. Do the same with spleen cells.
4. Spin Ficol gradient at 2500 RPM at 4 deg. C for 15 minutes.
5. Remove buffy coat by gently pipetting and transfer it to a fresh conical tube. Q.S. to 15 mL with IMDM. Add 10 µL to 10 µL trypan blue for cell counts. Pellet the cells at 1500 RPM. Carefully aspirate supernatant. Resupend pellet by flicking tube and resuspend in IMDM. Adjust volumes to 1 million cells/mL.

Purification of Lin-, c-kit+, Sca-1+ bone marrow cells for Culture, Flow Cytometry, or Transplantaion

M. Fero 6/05

Materials

Media

• Heat inactivated FBS (56°C x 30 min)
• PBS + 2% heat-inactivated FBS
• IMDM + 20% heat-inactivated FBS
• Viral transduction: Iscove's + 20% heat inactivated FBS, plus 100 ng/mL each of SCF, IL-6, and Flt-3L, plus 10 ng/mL IL-11.
• CFU assays: aMEM +30% FBS, 1% deionized BSA, 1.2% methylcellulose, 0.1mM ßME plus 3 U/mL Epo, 50 ng/mL Tpo, 100 ng/mL SCF, 100 ng/mL IL-6, ? ng/mL IL-3.

Antibodies for Lineage Depletion (Pharmingen)

Antibodies for Sorting (Pharmingen monoclonal rat anti-mouse)

Mice

C57 (Ly 5.2)
C57/SJL (Ly 5.1, Pep3b)

Other Reagents

Dynabeads (Sheep anti-rat, Dynal #110.08)
Ficoll
CH296
Methylcellulose (Fisher)
Deionized BSA (Sigma)
Protomine Sulfate

Cytokines (Peprotech)

mu SCF
hu IL-6
hu Flt-3L
hu IL-11
Thrombopoeitin
hu IL-3
Erythropoeitin (Amgen)

Procedure

Harvest Marrow

• Flush femurs with PBS/FBS and pass cells through a 23G needle.
• Also collect T-cells as a flow cytometry control.
• Collect low density cells by equilibrium centrifugation over Ficoll-Hypaque (d=1.077 g/mL).
  Alternatively, RBC may be lysed. See blood flow cytometry protocol for details.
• Wash cells in PBS/FBS.

Lineage Depletion

1. Resuspend at a density of 5x107 cells/mL in PBS/FBS plus antibody cocktail (each antibody final dilution = 1/500 v/v).
2. Incubate on ice for 30 min. Meanwhile, wash Dynabeads 2x in PBS/FBS. Resuspend the beads at their original concentration (4 x 108 beads/mL).
3. Wash cells in PBS/FBS, spin, and resuspend at 108 cells/mL.
4. Slowly add 1 vol. of Dynabeads (4 beads/cell) to the cell suspension. Incubate 5 min. @ room temp.
5. Expose to magnetic field in a 3 mL round bottom tube or eppendorf tube with the correct sized magnet. Transfer non-adherent cells to a new tube.
6. Remove tube with beads from magnet. Add 1/2 vol. PBS/FBS, gently mix. Expose to magnet again and pool non-adherent cells with the first.
7. Optional: Repeat the lineage depletion with a new round of antibody and beads (steps 1 - 5).
8. After the last mangetic bead depletion expose the cells to the magnet a second time to remove any remaining beads. Again, save only the non-adherent cells.

Cell Sorting (Lin-,cKit+,Sca1+)

1. Incubate cells in FcgRII block on ice for 10 min.
2. Stain an aliquot and with PE-IgG2a and FITC-IgG2b isotype antibodies as neg. controls. Incubate remainder of cells with Sca-1 PE, plus c-kit FITC antibodies on ice for 30 min (all antibodies 1/100 v/v).
3. Wash with 10 mL of PBS/FBS, spin and resuspend in PBS/FBS with 1 µg/mL of propidium iodide. Filter through 70 µm nylon mesh.
4. Select cells which are PI negative, c-kit positive, Sca-1 positive with intermediate forward and side-scatter. (Run T-cells as a size control).

Viral Transduction

1. Pre-coat 12 well plates with CH-296. To each well add 1.5 mL of Iscove's/20% FBS plus growth factors (SCF, IL-6, Flt-3L, IL-11).
2. Collect 5000 cells per well into 12 well plates with 1.5 mL/well of Iscove's transduction media.
3. Incubate 48 hrs at 37°C, 5% CO2.
4. Harvest cells by spinning at 1000 rpm for 5 min.
5. Add 0.5 mL of 3x growth factors to each well (depending on the experiment).
6. Resuspend cells in 1 mL of viral supernatant (or control media) and return to wells.
7. Incubate 12 hrs at 37°C.
8. Repeat viral transduction (steps 4 to 7). Incubate another 12 hrs at 37°C.
9. Harvest cells and replate in 1.5 mL of Iscove's/FBS plus growth factors (no virus).
10. Incubate 4 days at 37°C.

Transplant

1. Harvest fresh competing marrow from a C57/SJL (Ly 5.1) mouse. Pass through 23G needle and resuspend in Iscove's/FBS at 106 cells mL.
2. Harvest cells, wash with Iscove's/FBS.
3. Spin at 1400 rpm for 5 min. and resuspend in 1.5 mL. Record cell count. Add 1.1 mL of cells to 1.1 mL of competing marrow cells. Transplant into lethally irradiated (10 Gy, 60Co) recipients.
4. Of remaining cells save some for FACS analysis for GFP + lineage specific antibodies; the remainder can be used for CFU assays in methylcellulose.

CFU Assays

1. For CFU assays: Plate an average 20 cells in 35 mm dishes with 1 mL aMEM and 1.2% methylcellulose plus growth factors.
2. Incubate at 37°C in 5%CO2 (plus 5% O2 if available).
3. Analyze for erythroid colonies at 8 days (and confirm by Benzidine staining).
4. At 12 days analyze for granulocyte/macrophage colonies and large (>1.5mm) mixed colonies (erythroid, meg, and granulocytes confirmed by Giemsa staining).

GFP and Cell Surface Marker Immunostaining

1. Incubate cells with lineage specific antibodies as described for cell sorting, above. If live cells are not needed fixation with dilute formalin or paraformaldehyde is O.K. but alcohol based fixatives should be avoided because they allow GFP to leak out of cells.
2. For unconjugated antibodies, incubate with an anti-IgG2 PE-conjugated secondary antibody.
3. Wash and incubate with P.I. as for sorting. Analyze on fluorescent microscope or FACS machine.

M. Fero — 6/01
(after C Gorman, M. Calos)

Materials

• 2x HEPES: 8 g NaCl, 0.105g NaHPO4, 6.5 g HEPES, H2O to 500 mL, pH 6.95 to 7.10 (try a range)
• 2M CaCl2: store in aliquots at -20°C
• DNA 4 mg/mL

Procedure

(The volumes can be scaled up as necessary)

1. Let all solutions equilibrate to room temperature.
2. Feed cells on a 10 cm dish with 7 mL fresh media. They should not be more than 50% confluent.
3. Add 0.5 mL of 2x HEPES to a conical tube.
4. To a separate tube add 61 µL of 2M CaCl2, plus 4 - 10 µg (1 - 2.5 µL) DNA, and q.s. to 0.5 mL by adding (438 µL) H2O (or try TE pH8.0 ). Mix with a pipet and transfer dropwise to the 2x HEPES solution. Mix and then immediately sprinkle on top of the cultured cells. Swirl gently and return to the incubator overnight.
5. Change the media the following day.

M. Fero — 4/11/00, updated 4/25/03

Materials

Plastic tubes with anticoagulant: PBS + 2 µL anticoagulant for each 100 µL blood to be collected
Anticoagulant: 10% Na2EDTA, pH=7.4.
250 µL of fresh mouse blood in plastic tubes containing EDTA.
RBC lysis buffer (388 mM NH4Cl, 29.7 mM NaHCO3, 25 µM Na2EDTA)
20.75 g. NH4Cl
2.5 g. NaHCO3
0.093 g. Na2EDTA
1 L. H2O
Hemocytometer
Hema-3 Stain set (Fisher)

Comment

EDTA acts as an anticoagulant by chelating Ca++ ions and inhibiting activation of the clotting cascade. If viable cells are needed for culture or sensitive biochemical assays then heparin should be considered in place of EDTA.  The lysis buffer is a hypotonic solution that causes osmotic lysis of RBCs, but leaves WBC intact. It must be given at 19-20x the volume of blood to achieve lysis.  Extra lysis buffer may be necessary if full RBC lysis is not achieved. An excess of lysis buffer generally will not hurt the WBC. If the WBC need further manipulation after RBC lysis, then wash the cells by centrifugation and resuspension in PBS or media.

Hematocrit (Packed red cell volume)

Draw blood in 2 heparinized hematocrit capillary tubes.
Spin 5 min in hematocrit centrifuge. Measure total and packed cell volume.
Calculate packed cell volume as a percent of the total.

Red cell count

Dilute cells 1/1000 in PBS.
Add 10 µL to a hemocytometer. Count the number of RBC per large square.
Calculate: RBC/large square x 1,000 dilution x 10 large squares/µL = RBC/µL blood.

White cell count

Add 10 µL whole blood to 190 µL of lysing reagent (a 1/20 dilution). Mix and incubate 1 min.
Add 10 µL of lysed blood to hemocytometer. Count the number of WBC per large square.
Calculate: WBC/large square x 20 dilution x 10 large squares/µL = WBC / µL blood.

White cell differential count

Spot 20 µL of whole blood near the frosted end of a glass slide.
Smear the drop out across the slide with the end of a second glass slide to obtain a thin film with a smooth feathered edge. Air dry the slide.

Hemastain: 5 dips in fixative, blot dry. 3-5 dips in Solution I, blot dry. 3-5 dips in Solution II, blot dry. Rinse 1 min by placing in a coplin jar under gently running dionized water. Air dry.
Under bright field oil microscopy assess the RBC morphology and perform a differential count on a total of 200 WBC.

Automated cell counts: For automated RBC, WBC and platelet total counts send 0.5 mL of blood (or blood diluted in PBS) in a purple top EDTA tube to the hematology lab. Send specimens before 3 p.m. and call ahead of time. A WBC differential requires more blood.

(PCR Recipe: Kogan et al, New Engl J Med 317: 985-990)

Primer Sequences

p27KO mice. JAX Stock 2781 (C57BL/6), JAX Stock 3122 (129S4) Fero ML, et al. Cell. (1996) 85:733-44
KO (knockout allele; Contains Neo)
Neo-1 (CCTTCTATCGCCTTCTTG)
K-3 (TGGAACCCTGTGCCATCTCTAT)
Size: KO(-) = 0.5 kB

WT (wildtype allele)
K-3 (above)
K-5 (GAGCAGACGCCCAAGAAGC)
Size: WT(+) = 1 kB

p27LoxP mice: Cre-inducible knockout (Chien WM, et al. PNAS (2006) 103:4122-7)
WT or L+ (exon 1 and 2 flanked by LoxP)
K-52 (TAGGGGAAATGGATAGTAGATGTTAGGACC)
K-53 (GGTATAATACGGAAAGTGACTCTAATGGCC)
Sizes: WT(+) = 500 bp, (L+) = 550 bp

L- (null allele, minus exons 1 and 2, residual LoxP, no Neo)
K-53 (GGTATAATACGGAAAGTGACTCTAATGGCC)
K-57 (AGCGGCTCCCGGCGCCGAGAC)
Size: (L-) 250 bp

p27STOP: Cre-inducible knock-on mice (Chien WM, et al. PNAS (2006) 103:4122-7)
WT and S+ (induced wildtype; contains residual LoxP site)
K-55 (CGCCTGGCTCTGCTCCATTTGAC)
K-56 (GACACTCTCACGTTTGACATCTTCC)
Sizes: WT(+) = 192 bp, (S+) = 240 bp

S- (Uninduced null allele; contains Lox-Stop-Lox)
K-55 (CGCCTGGCTCTGCTCCATTTGAC)
Neo-5 (CTACCCGCTTCCATTGCTCAG)
Size: (S-) = 431 bp

Buffer Recipes

Stock Solutions
1 M Tris pH 8.8 (do not use pH meter, store at room temp.)
1.23 g Tris HCl
5.13 g Tris base
q.s. 50 mL with H2O

KG-1 (10x)
8.3 mL 1M (NH4)2SO4 [10x = 166mM]
33.5 mL 1M Tris base pH 8.8 [670 mM]
174 µL ß-Mercaptoethanol [50 mM]
3.35 mL 1M MgCl2 [67 mM]
q.s. to 50 mL with H2O, and aliquot into 1.5 mL Eppendorf tubes
Store Frozen

KG-2 (10x)
25 µL of 100mM dNTP Stocks (x4) [@10 mM]
25 µL DMSO [10%]
25 µL 8 mg/mL BSA [0.8 mg/mL]
150 µL H2O (makes 250 µL total vol.)
Store frozen
PCR Master Mix (Multiply volumes x # of PCR reactions, below)
2 µL KG-1 (10x) [1x final]
2 µL KG-2 (10x) [1x]
2 µL Primer 1 (1uM) [0.1 mM]
2 µL Primer 2 (1uM) [0.1 mM]
0.2 µL Taq (Gibco) (5 U/uL) [0.05 U/mL]
10.8 µL H2O (subtotal 19 µL)
Aliquot into PCR tubes (19 µL each)
Add 1 µL sample DNA to each tube (20 µL total vol.)

Cycling Parameters

Do not place the tubes on the machine until the block has heated to > 90ºC. The first 4 cycles use a higher melting temperature to help denature long genomic DNA. Subsequent cycles use a lower melt temp, which is sufficient to denature shorter PCR products and preserves enzyme function. Longer (2 min.) extension times should be used if products > 2 kb are being amplified.
95°C hold x 2 min. (insert tubes at > 90ºC)
(96°C x 30", 57°C x 30", 65°C x 1-2') x4
(93°C x 30", 57°C x 30", 65°C x 1-2') x36
4°C hold

Run products on a 0.8-1.2% agarose gel

(M.Fero 10/2013)

Reagents

Lysis Buffer: (10 mM Tris pH8, 100 mM NaCl, 25 mM EDTA, 0.5%SDS), store at room temp.

10 mg/mL Proteinase K: 100 mg (Roche) + 10 mL buffer (10 mM Tris pH8.0, 20 mM CaCl2, 50% (v/v) glycerol), store at -20ºC in 1.5 mL aliquots.

Lysis buffer + proteinase K (1 mg/mL): 50 mL Lysis buffer (above) + 0.5 mL 10 mg/mL proteinase K, store at -20ºC.

TE: 10 mM Tris pH8, 1mM EDTA

Phenol: Molecular biology grade. Equillibrate 1x with Tris pH8, and then 1x with TE Store at 4°C.

Chloroform: Fresh choloroform + 1/20 vol. isoamyl alcohol. Store at r.t.
Absolute ethanol.

Procedure

1. Cut 3 mm of toe or tail or toe tips from 7-10 day old mice into 1.5 mL microcentrifuge tubes.
2. Digest tail biopsy by adding 0.7 mL lysis buffer + proteinase K and incubate for 4-16 hrs at 37°C.
3. Add 0.7 mL of phenol (or a 1:1 (v:v) mix of chloroform:phenol) and slowly rotate at 4°C for 1 hr. to overnight.
4. Spin at high speed (15,000 g) for 2 minutes to separate phases. The hair and other debris should pellet to the bottom. Pipet the top (aqueous) phase to a fresh tube, taking care to avoid the bottom (organic) layer.
5. Repeat the extraction (as in steps 2-3) with 0.7 mL of phenol or a 1:1 (v:v) mix of chloroform:phenol for 1 hr.
6. Repeat the extraction (as in steps 2) but with 100% chloroform. You should now have ~0.7 mL of aqueous solution in each tube. Spin for 2 min. Transfer 0.5 mL the solution to a fresh tube, taking care not to place the pipet tip to the bottom of the tube (since this is where any residual CHCl3 will be located).
7. Precipitate by adding 2 volumes (1.0 mL) of absolute ethanol and mix vigorously. You should see some DNA stranding at this point. Incubate at -20°C for >30 min. Samples may be stored in -20ºC in ethanol indefinitely.
8. Spin at 12,000 rpm for 5 min. Discard supernatant. Wash pellet by adding 1 mL of 70% ethanol, mix by inversion, and spin for 1 minute. Discard supernatant being taking care not to pour out the pellet. It is likely to be loosely adherent since the salt is washed out. Repeat the 70% ethanol wash, spin and again discard the supernatant. Blot the excess ethanol carefully on a clean paper towel. Invert the tube on a paper towel to air dry the tube.
9. Resuspend pellet in 0.1-0.5 mL TE (depending on the amount of DNA). Use 1 µL for a PCR reaction, or 1/2 of the sample for a Southern blot. Store the genomic DNA at 4ºC for 1-2 weeks, or at -20ºC for longer periods.

Quality Control - Options for QC include the following:

1. Run 2 µL of the genomic DNA on a 0.6% agarose gel, with high MW marker, e.g. lambda HindIII. The DNA should form a high MW band (20-50 kb), with minimal smearing or degradation below this level. RNA may also be present if a metabolically active source tissue was used instead of toe/tail tip. Insoluable material, or protein contamination, may result in ethidium bromide staining that does not migrate out of the loading well.
2. A spectrophotometer (e.g.Nanodrop) may also be used to measure the A250/A280 ratio, as a crude indicator of DNA purity. Values of 1.9-2.0 are expected. Values > 2.0 may indicate organic solvent contamination; values <1.9 represent protein contammination.

Quantitation - Options for quantifying genomic DNA include:

1. The DNA concentration may be estimated by comparing band intensity on an agarose gel to lamda HindIII or other standards with known band quantitities.
2. The spectrophotometric A260 value may be used to estimate DNA concentration, if the DNA is high quality. ([DNA] mg/mL = A260 x 50)
3. Fluorometry with a DNA binding dye (SybrGreen or Höchst) is an accurate and sensitive measure of DNA concentration.

What is the MPD?

The MPD is a laboratory relational database that allows users to cross-reference mouse inventory data, breeding records, pedigrees, pathology data, histology images, autoradiographs, PCR genotyping protocols, freezer archives, and reagents (specifically plasmids, oligonucleotides, and antibodies). It is comprised of a set of FileMaker templates which can be used on a single computer or on a server with multiple clients. The Database is modular with the major components in separate files so undesired parts can simply be discarded with only minor modifications to the Directory (to prevent error messages).

Download the install and initial setup instructions.

M. Fero 1/8/09

Procedure

1. Register online
2. Bring mouse in clean cage to imager
3. Sign in on log sheet
4. Clean inside of Xenogen system and external mouse anesthesia chamber. To minimize background, use black plastic background sheet for fluorescence, use black paper for luminescence.
5. Anesthesia
   1. Turn on gas cylinder (but don't change gas regulator on cylinder)
   2. On Xenogen box - turn 'Gas' knob to 'on' (always leave power button on).
      1. On anesthesia machine:
         1. O2: On
         2. Vaccuum: On
         3. Switch up (open) valves to Chamber and IVIS. Set flow rate with ball valve to midway (~2 L/min).
         4. Set guage on lid of Isofluorane container to '2' (can change to alter level of anesthesia).
         5. Place mice in chamber.
6. Login to computer
7. Living Image #3 software
   1. Launch program (icon on Desktop). Enter your initials to tag images.
   2. Control Panel: Initialize IVIS System
   3. Choose field of view (A-D) depending on # of mice to image simultaneously.
8. Position animals: (Alternatively the plastic phantom mouse can be used with a fluorescent insert).
   1. Open door: Green laser rectangle should appear
   2. Place nose cones / corks in manifold and position at back of green rectangle
   3. Transfer mice from external chamber to IVIS nose cones. Turn off gas valve to chamber.
9. Acquire images:
   1. IVIS Acquisition Control Panel
      1. Select Fluorescence vs. Luminescence
      2. Exposure time: default = 1 sec.
      3. Binning: default = medium (4 pixels per bin). Decrease if saturated, increase if very faint.
      4. f/stop: default = 2
      5. Excitation/Emission: Set according to fluorochrome. GFP/FITC = preset #3? (see phantom mouse booklet)
      6. Select 'Acquire Image'
   2. Image Window:
      1. Counts should be between 600 and 600k? (or 60k?)
      2. Change units from Counts to Efficiency (to compensate for increased efficiency at center of image)
   3. Multi-sequence: In Control Panel select 'Sequence setup'.
      1. Set delay time, wavelength, exposure, etc.
      2. Acquire images
      3. In image window select 'display all' icon (upper right) to manipulate individual images.
10. Image analysis (can be done post hoc with Living Image software in computer facility)
    1. Set ROI and filters to quantify regions of interest.
11. Save images to desktop in Fero Lab folder. Map Fred network drive and transfer your files. Then disconnect Fred.
12. Cleanup
    1. Return mice to cage
    2. Turn off Isofluorane. Flush for 1 min. then turn off gas flow on anesthesia machine and gas cylinder (opposite of 5.3, above).
    3. Use a paper towel moistened with Clidox to clean all surfaces that have been in contact with mice.
       1. Inside Xenogen chamber, Plastic backing, nose cones, door handle.
       2. External anesthesia chamber.
          Area around mouse cage.
13. Sign out on log sheet. Return mice to their proper room and racks.

<R. Gu 4/6/2005

Materials

24 well dishes
Millicell 0.4 µm inserts (Millipore PICM01250) - coated with fibronectin
Culture medium
PBS
Fixative: 4% paraformaldehyde in PBS (pH 7.6)
Rabbit anti-GFP (Molecular Probes, A-11122)
FITC-goat anti-rabbit IgG (Jackson ImmunoResearch, 111-096-006)
Biotinylated anti-BrdU (AbCam, ab2284-125)
Streptavidin-Alexa Flour 555 (Invitrogen/Molecular Probes, S21381)
Goat serum
Horse serum
BSA
2N HCl
DAPI Fluorescent mounting medium (Vectashield, Vector Labs H-1200)
Glass microscope slides and cover slips

Procedure

1. Coat Millicell inserts with fibronectin as described by Millipore.
2. Cultivate tissue at 37ºC on fibronectin coated well inserts in a 24-well dish.
3. Remove well inserts and cut out mesh insert with a scalpel, forceps and dissecting microscope
4. Place inserts in a 2.0 mL Eppendorf tube with 1 mL of blocking agent (10% goat serum, 1% BSA in PBS). Gently flick tube to mix.
5. Remove solution and wash 3x by adding 1 mL PBS, 3' each.
6. Add 0.3 mL rabbit anti-GFP (1:200 v/v, plus 1% goat serum in PBS) and incubate 1 hr. at room temp. Wash 3x in PBS.
7. Post-fix in 1 mL 4% paraformaldehyde x15', room temp. Wash 3x in PBS.
8. Denature DNA with 1 mL of 2 N HCl x30'. Wash 3x in PBS.
9. Block in 1 mL 10% Horse serum, x30' at room temp. Wash 1x in PBS.
10. Add 0.3 mL Biotinylated anti-BrdU (1:300, with 1% horse serum in PBS). Incubate at 4ºC o.n. or 2 hr. at room temp. Wash 3x in PBS.
11. Dilute 1 µL fresh streptavidin-Alexa Flour 555 in 1 mL PBS (1:1,000). Spin the dilute antibody at high speed for 30' to remove particulates. Add supernatant to specimen and incubate 30' at room temp.
12. Remove antibody and add 1 mL PBS. O.K. to store at 4ºC.
13. Use a scalpel to remove membrane from well inserts. Spot 25-50 µL of DAPI mounting medium onto a glass slide. Place 1-2 membranes on the DAPI medium. Use a dissecting microscope to be sure to place the tissue side up. Cover with a 24-40 mm glass slip. Paint the edges with clear nail polish. View with immunoflourescence. Store in the dark at 4ºC.

M. Fero

Materials

Syringes, 1 mL with 36G needles
BrdU: 3-10 mg/mL in PBS
Fixative: Methyl Carnoy's, or 4% paraformaldehyde in PBS
Paraffin processing materials
Organic solvents: Xylene, EtOH, MeOH
0.3% H2O2: 2 mL 30% H2O2 in 200 mL MeOH
Trypsin: Dilute (10x stock, -20ºC) 1/5 in PBS
2.5 N HCl: Dilute concentrated HCl 1/22 (v/v) in H2O
Mouse anti-BrDU: DAKO cat#M0744, $205/mL
Biotinylated horse anti-mouse IgG, Avidin, biotin-HRP, Horse serum: Vector, Vectastain ABC kit, #PK4002 $185
Color substrate, (make fresh): 25 mL 50 mM Tris pH8, 1 mL DAB (3,3-diaminobenzidine, 25 mg/mL stock, -20ºC), 100 µL NiCl2 (10% w/v stock, 4ºC), 10 µL 30% H2O2.
Counter stains: Methyl Green or dilute Eosin (1/4x in 80% EtOH)

BrdU Labeling

1. Inject 10 µL of BrdU per gm of body weight (30-100 mg/kg). Use higher doses for short labeling times. Label for 1-2 hours to measure the number of cells in S-phase. Maximal staining intensity will occurr if the labeling time exceeds the duration of S-phase (6 hours) Longer labeling (e.g. 1-7 days) does not give the number of cells in S-phase at a given time point but will give the proliferative fraction for that time period. To label longer than 12 hours give repeated doses every 24 hours. Use lower BrdU dose (30 mg/kg) for prolonged labelling to avoid chemotherapy-like toxcity. Lower doses of BrdU also effects the intensity of staining and indirectly the number of cells that will be deemed "positive".
2. Control: The best negative control is an additional animal given no BrdU (or more formally - injected with PBS only). This is a "no antigen" control and is superior to using "no primary antibody" as a negative control to accurately judge the level of background staining. Use small intestine as a positive control since this gives robust labeling in the epithelial crypts even with short labeling times.

Tissue Preparation

1. Place freshly dissected tissues in tissue cassettes and fix either in Methyl Carnoy's for 1 hour or paraformaldehyde overnight.
2. Embed in paraffin, and cut 4 µM sections on glass slides (see Tissue Processing protocol).

Immunostaining
In Plastic Tubs (200 mL):

3. Deparaffinize:
   Xylene 2' (x3)
   100% EtOH 2' (x3)
   70% EtOH 2' (x1)
4. Quench endogenous peroxidase activity (optional):
   MeOH 2' (x1)
   0.3% H2O2 in MeOH 20' (x1)
5. PBS 5' (x3).

In Coplin jars (25 mL):

1. To decrease antigen masking by chromatin proteins digest in 2X Trypsin (1mg/mL in PBS) 10' (x1) and then wash in PBS 2' (x3)
2. Denature (depurinate) the DNA with 2.5 N HCl, 37°C 15' (x1), wash in PBS 2' (x3). (0.1N NaOH can also be used but may be harsh on certain tissues). Steps 6 and 7 may need to be altered if they destroy other antigens necessary for double immunostaining.

In Humid Chambers (at each step use 100 µL and cover with glass slips. To remove the slips dip the slide in a jar with PBS. Don't pull off the slips as this will scratch the specimen):

1. Block with Horse serum (1/80 in PBS) 30' (x1). Wash in PBS 2' (x3)
2. Primary antibody 100 µL (1/200 mouse a-BrDU) (r.t. or 37ºC) 1 hr (x1). Wash (x3).
3. Secondary antibody (biotinylated horse anti-mouse IgG) 1/250, 30' (x1). Wash (x3)
4. 100 µL Avidin + Biotin-HRP mix (1/125 v/v Sol.A, 1/125 Sol B), 30' (x1). Wash (x3)
   In Coplin jars (25 mL):
5. 50 mM Tris pH8 37°C (x1)
   DAB/NiCl2 Color substrate 37°C, 3' (x1).
   Rinse under H2O tap for 5 minutes.

In Plastic tubs (200 mL):

1. Counter stain in methyl green for 20' or dilute Eosin (1/4x in EtOH) for 2'.
2. Dehydrate in graded ethanol:
   95% EtOH 1' (x2),
   100% EtOH 1' (x2),
   Histoclear or Xylene 1' (x2) (Methyl green is not stable in xylene)
3. Add 1-2 drops of Permount with a glass rod and cover with glass slip. Cure overnight in fume hood. Don't stack slides on top of each other until fully cured (2 days).

Euthanasia

Euthanasia and mouse necropsies require prior IACUC approval. The mode of euthanasia should be chosen which minimizes pain or distress to the animal and provides biologically meaningful samples. Only personnel with appropriate animal training may handle live animals.

Isoflourane inhalation: This should be given in sufficient dose to cause complete loss of breathing and pulse in the mouse. Apply 1 mL to a gauze pad in a bell jar. A porous barrier should be placed over the guaze pad so the mouse does not come in direct contact with the anesthetic. (Note: Halothane inhalation in humans has been associated on rare occasions with hepatitis. In the laboratory setting this risk is infintesimely small but its use nonetheless has been discouraged). Use gloves and an externally vented fume hood for personal protection.

Carbon Dioxide inhalation: CO2 is an effective agent for euthanasia and may be used if a suitable CO2 source, and container is available. For histological purposes it has the disadvantage that it induces capillary hemorrhages in certain tissues, particularly the lungs and brain. Dry ice should not be used because it can cause frostbite in the animal and is does not provide a controlled dose of carbon dioxide. Sufficient quantities of CO2 should be used to induce rapid loss of consciousness and be continued to respiratory motion ceases. CO2 induces a metabolic acidosis and suboptimal levels may induce agitation and hyperventillation in animals. For this reason its use has been disparaged but in practice there is little behavioral evidence of distress, when used appropriately, and it does not inferior in this respect to expensive anesthetics.

Cervical dislocation: This should only be performed on anesthetized animals which are unresponsive to noxious stimuli, such as a foot pinch. It is the most quick, effective and humane way to ensure the death of the animal. It is useful to ensure that anesthetic clearance does not lead to partial recovery. (Inhaled anesthetics may decrease due to tissue redistribution or respiratory activity). Because it may introduce traumatic artifact it should not be performed in the case where CNS or pituitary hisology is needed. Cervical dislocation of unanesthetized animals should only be performed by experienced personnel if necessary for experimental purposes and requires IACUC approval.

Fixation

Fresh tissues should be immediately placed in plastic histology cassettes and immersed in an appropriate fixative to avoid tissue autolysis. Intestine, liver, prostate and brain are particularly prone to autolysis. Other organs can be kept a short time in cold PBS. After fixation the tissues should be transferred to 70% ethanol and stored at 4°C until processed. If RNA is to be purified the tissues should be harvested quickly, placed into eppendorf tubes or plastic vials and snap frozen in liquid nitrogen.

Formalin: Formalin solution, 10% neutral buffered (Sigma). (This contains 4% w/v formaldehyde with phosphate buffers) Formalin stabilizes proteins cross links proteins and prevents decomposition. The solution may be used several until it begins to become discolored. It is good for paraffin sections and H&E stains. Tissues should be fixed overnight, or less if they are very thin (less than 2 mm).

4% paraformaldehyde: Paraformaldeyde is the anhydrous form of formaldehyde. It needs to be made fresh in PBS or stored frozen. It is slightly less harsh than formalin and is sometimes preferred for immunostaining.

Bouin's: This fixative comes prepared from Sigma and is sometimes preferred for immunostaining with certain antibodies. It's yellow color causes can cause some undesirable discoloration of tissues.

Methyl Carnoy's: (10% glacial acetic acid, 60% methanol, 30% chloroform) This is a rapid fixative which requires only a few hours to completely permeate and fix tisues. It is routinely used by pathology lab for immunohistochemistry, but like all fixatives its utility for this purpose is antigen and antibody specific. H&E stained tissue will appear very eosinophilic (reddish) in this and other alcohol based fixatives.

OCT: This clear jelly is not a fixative but is the embedding compound for frozen sections. Frozen sections work best for immunostaining but it is difficult to cut thin frozen sections and get as good of morphology as with paraffin. Tissues can be quick frozen by placing them in 1.5 mL tubes and plunging this in liquid nitrogen prior to storage at -70°C. Alternatively frozen sections can be prepared immediately by placing tissues in molds with OCT.

B5: B5 is a mercury based fixative that provides superb morphology for hematopoietic cells. Mercury is a hazardous chemical so this fixative should be used of sparingly and must be disposed of with EHS approval in a specially designated container.

Dissection

Record the identifying data plus the appearance and behavior of the animal. The general approach is to record the weight of the animal, each of the individual organs and the carcass after the organs are removed. If skeletal defects are suspected save the carcass and obtain a radiograph of the skeleton. Animals should be laid upon a clean paper towel and have all 4 extremities pinned to thin styrofoam or cork board.

Body Wall: Wet the animals fur with 95% ethanol to minimize contamination with hair. Instruments should be soaked in an ethanol if cultures are to be obtained from tissues. Dedicated scissors should be used on skin and bone. (Wash blood and tissue from instruments before soaking them in ethanol or flaming them. Otherwise the tissue will become fixed to the instruments)

Skin: A vertical, ventral, midline incision should be cut with scissors from the neck to pubis. This should be extended laterally at both axillae and inguinal areas to form an "I" shape. Reflect the skin laterally and pin down. Abdominal skin samples should be saved as vertical thin strips (3 x 15 mm) cut with a scalpel (In this way sections will be parallel to the hair follicles)

Mammary tissue: The inguinal and axillary mammary glands are visible as yellow fat pads adherent to the underside of the skin beneath a fascial plane. They may be separatd from the skin by grasping them with forceps and sharply dissecting (i.e. with a scalpel) them away from the underlying skin with a gentle stroking motion.

Abdomen: Fresh sterile scissors should be used to open the peritoneum if cultures of MEFs are to be made. In this case pin the body wall back with the skin.

Uterus and ovary: A gravid uterus is appaarent by it's "beads on a string" appearance in the lower abdomen. The murine cervix has two horns which extend up from a cervix in the pelvis. Cut across the cervix and lift out the uterus with forceps. As the uterus is lifted fat and myometrial attachments may be separated. At the superior end of each uterine horn lies an oviduct and an ovary. The ovaries are located just below each kidney with which they share common blood vessels. Following fixation the uterus should be cut into thin cross sections with a scalpel prior to being processed into paraffin. The ovaries should be wrapped in lens paper and embedded in a block with other small organs such as the adrenal, thyroid, lymph nodes and pituitary.

Kidney: The kidney should be cut lengthwise with a scalpel through the renal pelvis prior to formalin fixation. The cut surface should be embedded face down for sectioning.

Liver: The 3 major lobes of the liver should be separated. 3-4 mm wide strips should be cut with scissors or a scalpel prior to formalin fixation. The cut surface should be embedded face down prior to sectioning.

Adrenal: This small (2mm) pyramid shaped organ looks like a little triangle of pinkish fat on the superior pole of each kidney. It should be wrapped in lens paper and placed in cassettes along with ovaries, pituitary, thyroid, or other small sections to avoid losing them during fixation and paraffin processing.

Spleen: The spleen should be cut lengthwise with a scalpel prior to formalin fixation. Small white specks (lymphoid follicles) can be seen. These may be counted on the surface of the spleen (which is the basis of the CFU-S hematopoietic assay). The cut surface should then be embedded face down prior to sectioning.

Pancreas: Pancreas is a fatty looking tissue adherent to the first part of the intestine on one end, and on the spleen on the other. It can be fixed whole.

Intestine: The intestine should be dissected clean from all mesenteric fat. The luminal contents should be gently squeezed out with the back of a pair of scissors or similar blunt object. It should be kept moist with saline and fixed quickly to avoid degeneration of the villi. Beta-gal staining is best accomplished by opening the intestine lengthwise and pinning the edges at 2 cm intervals down on a wax lined tray. After staining the intestine can be folded in pleats and placed in cassettes for formalin fixation. For longitudinal sections, cut the folded intestine lengthwise and embed with the cut surface down. Cross sections are more difficult to obtain but one method is as described for uterus.

Prostate and Testes: The testes are easily retrieved from an open abdomen by traction on the spermatic cord. They should not be cut prior to fixation because they will rupture, instead fix them whole. The prosate, in the mouse, contains several different lobes. The anterior prostate is a mat of soft tissue which lies tucked into proximal fold of the large, white, convoluted, seminal vessicles. Take care when handling the seminal vessicles or the spermatic fluid will make a mess of the dissection. They may be trimmed near their attachment to the proximal urethra. At the base of the bladder the seminal vessicles, ductus deferens converge onto the proximal urethra. Situated circumferentially on the proxmal urethra are a series of soft tissue protuberances. Posteriorly, the dorsal lobe forms a pair of wing like structures which may be fused with a small protuberance more laterally, together forming the dorsolateral prostate. On the anterior side of the urethra the ventral prostate forms one or two small bumps. The neck of the bladder, proximal urethra and prostatic lobes may all be fixed en bloc which will facilitate proper orientation during embedding. Alternatively the individual lobes may be separated. If separated lobes are fixed in one cassette it is useful to mark them with dissecting ink to facilitate their identification in histological sections.

Thorax: The thorax should be opened by cutting away the rib cage with scissors. The thymus or pericardial fat may be adherent to the inner chest wall.

Thymus: The thymus has two lobes and sits on the superior and ventral aspect of the heart. It involutes in older animals and becomes increasingly surrounded by fat (older than 3 months). Under the dissecting microscope the texture of thymus can be distinguished from fat and large vessels more easily. If lacerated the contents of the thymus may spill so it should be fixed whole. Each lobe of the thymus should be embedded with the superior edge down. If frozen sections are to be performed then molds for OCT may be formed from the back end of a VWR marker. Insert a single lobe into the OCT superior surface down and place on a dry ice + ethanol bath. Be very careful not to get any ethanol into the OCT or it will turn the frozen block into a slimy mess.

Heart: The heart should be cut in cross section with a scalpel in 3 mm strips.This will create a set of "doughnuts" which should be fixed and then embedded cut surface down.Both the right and left ventricles will be visualized in this fashion.The upper heart should also be sectioned, this consists of cardiac valves, atria and the large vessels.

Lung: The lung can be cut into strips or fixed whole. Don't squeeze air out of it, that makes it harder see the normal alveoli.

Lymph nodes: Lymphomas will cause generalized enlargement of lymph nodes, spleen or thymus. Be aware that infections or skin ulcers will cause enlargement of the nearby nodes and may need histology be distinguished from lymphomas. Infections or severe anemia may also cause splenomegaly. Lymph nodes are small (1-2 mm) pale tan ovoid structures to be found in the axilla, inguinal area, and in the abdomen. They should be fixed whole. Lymphomas can be sectioned prior to fixation and embedded cut surface down. Small lymph nodes in normal areas are found in these same areas but are difficult to identify without a dissecting microscope. Like all small organs they should be wrapped in lens paper to avoid loss during fixation.

Extremities

Skeletal muscle: This can be cut from the thigh bones and fixed whole.

Bone marrow: The entire femur from the hip joint to the knee should be cut away from the animal with scissors. All of the muscle should carefully be cut away. Cut off each end of the bone with scissors to reveal the hollow marrow filled core. Bone should be fixed whole in formalin for 24 hours. It then should be transferred to decalcification (11% formic acid with stirring) solution for overnight prior to paraffin processing. One femur should be saved for cytology as well. With a 26G needle flush 1 mL of media (Iscove's +15% heat inactivated serum) through the marrow space and out the other end of the bone into a 1.5 mL tube. Plate 100 µL onto positively charged glass slides with a cytospin centrifuge and stain with Wright-Giemsa. This will preserve the cellular morphology and will allow an accurate differential cell count to be performed.

Head and Neck

Mouth: Inspect the mouth for excessive growth of the incisor teeth or abnormal jaw structure. Excessive incisor growth can prevent mice from feeding normally and can cause malnutrition.

Thyroid: The strap muscles connecting running along the neck should be cut away with scissors under a dissecting microscope. When the trachea is well exposed it should be grasped inferiorly with forceps and retracted downwards. It then should be cut above the larynx several mm below to remove it from the neck. The thyroid gland has the same color as muscle and consists of two small patches of tissue adherent to the trachea just below the laryngeal prominence. Under the stereomicroscope it will appear more translucent than muscle. Do not remove the thyroid from the trachea but instead cut across the trachea just above and below the thyroid tissue. For better morphology fix and embed this en bloc and cut cross sections across the trachea.

Eyes: The eyes will protrude slightly if the fur is retracted. Grasp the orbits with forceps and cut them free from the ocular muscles and optic nerve. For optimal retinal morphology a sharp slit in the corneas should be made and the lenses removed. To accomplish this make a hole in the cornea with a needle and the cut away the cornea with very fine scissors. Gently remove the lens using fine forceps without distorting the remainder of the eye or detaching the retina. After the corneas and lenses have been removed little hollow cups will remain. Fix these in formalin for paraffin or 5% glutaraldehyde for plastic embedding. Embed the retinal cups on their sides. When sectioning cut the blocks down to the level of the optic nerve for optimal morphology.

Brain: If optimal CNS morphology is required consider infusing fixative intravascularly via intracardiac perfusion prior to dissection. Retract the skin from the skull. The cranial bone should be excised along its perimeter. Start by flexing the neck and inserting the tips of scissors into the foramen magnum at the skull base. Cut the cranium with small snips at its juncture with the facial bones, superiorly to the reveal the entire brain and olfactory lobes. Gently lift the brain under the frontal lobes and free it from underlying vessels and cranial nerves. Remove in one piece the cerebrum, cerebellum and as much of the hind brain as possible. Do not cut the brain until it has been fixed. After paraffin embedding the brain should be cut with a scalpel and embedded cut surface down. At this time you must decide whether you want to view coronal or sagital sections.

Pituitary: After the brain is removed the pituitary will generally remain sitting on the floor of the skull between the two large trigeminal nerves that run longitudinally along the bone. The pituitary is a 2 x 4 mm strip of pale tissue running crosswise between these two nerves. Under the dissecting microscope the thin meningeal membrane that overlies the pituitary should be torn away with forceps. Gently lift out the pituitary with the tip of a curved 26G needle or with fine forceps. Let it sit in a drop of fixative for a few minutes and then wrap it in a small piece of lens paper and transfer it to a tissue cassette for fixation with other small organs. All 3 tissue layers will be visualized if it is embedded upside (posterior) side down or if cut in the coronal plane.

Cleanup

Although laboratory animals are unlikely to carry human pathogens yet tissues should be treated as potentially infectious and disposed of in biohazardous waste. Mice harboring human cell or tissue xenografts have a greater potential to carry human pathogens, and should be handled with greater precautions according to Biosafety Level (BSL) 3, otherwise mouse tissues are considered BSL 2. Carcasses should be frozen in plastic bags and disposed of the designated location in the animal facility. Return cages to the cage wash area in the animal facility, but refrain from entering animal housing areas after handling dirty cages to minimize the risk of pathogen transmission.

M. Fero 4/6/2006
(courtesy of Katie Rudd, Trask lab)

Materials

20 ng BAC DNA clone grown o.n. in 5 mL LB + chloramphenicol
Autogen reagents
Slide warmer
Organics: MeOH, EtOH, formamide
Fixative (3:1 MeOH:glacial HOAc)
Biotin (or digoxigenin) nick-translation Kit (Invitrogen)
20x SSC (3 M NaCl, 0.3 M NaCitrate, pH 7.0)
TE, pH8
CoT DNA 1 μg/μL (Invitrogen)
Glass slides, 18 mm glass slips
Rubber cement
3 M NaOAc
Na Azide solution (e.g. 25% stock)
Nonfat dried milk
Formamide
CHAPS or TWEEN detergent
Streptavidin-FITC (Vector)
Biotinylated goat anti-streptavidin (Vector)

Procedure

Day 1: Prepare cells and DNA

1. DNA:
   1. Prepare BAC DNA probe on Autogen machine (see separate protocol)
   2. Run 4-6 Autogen tubes per BAC.
   3. Hyb buffer: 50% formamide, 2x SSC, 10% dextran sulfate
2. Cells: Feed control cells with fresh RPMI + 15% BSA.

Day 2: Check DNA and spots cells onto slides

1. DNA:
   1. Resuspend DNA in 50 μL/autogen tube and pool in a 1.5 mL tube.
   2. Digest with Eco RI (5 μL DNA, + 1 μL enzyme, + 1.5 μL 10x buffer, + 7.5 μL H2O) 37ºC x 1-2 hrs.
   3. Run on 0.8% agarose gel (in 0.5x TBE gel + 0.5 μg/mL ethidium Br) x 2 hrs, to check DNA quality.
2. Cells:
   1. Pellet cells and wash 1x in PBS.
   2. Optional: Treat cells with hyptonic solution.
   3. Pellet and resuspend in 3:1 MeOH:Glacial Acetic acid
   4. Cells can be kept at -20ºC in fixative, but if so they should be washed with fresh fix on the day of use.
   5. Place a wet paper towel on 37ºC hot plate.
   6. Wash slides by placing in 100% MeOH in Coplin jar at room temp.
   7. Wipe both sides with Kimwipe
   8. Breathe on slide to moisten surface
   9. Drop two spots of cells from a Pasteru pipette 6 - 12'' above slide.
   10. Place slide on moist paper towel on warmer for 1 min.
   11. Move to dry part of slide warmer and leave it here o.n. to age (or place in 65ºC oven x 1hr.)

Day 3: Label DNA and setup hybridization

1. Setup nick-translation:
   1. 15 μL BAC DNA (~ 1 μg)
   2. 5 μL 10x dNTP mix
   3. 5 μL enzyme
   4. 25 μL H2O
   5. Incubate at 16ºC x2 hrs, then place on ice.
   6. Run a 5 μL aliquot on a 1% agarose gel to confirm fragmentation in the 200-600 bp. (If fragments are too large the DNA then return tube to 16ºC for 1 more hr.)
   7. Stop reaction with EDTA solution. (DO THIS BEFORE ADDING CoT!)
2. Add 1 μL CoT DNA (1 μg)
3. Ethanol precipitate by adding:
   1. 200 μL TE (final vol 250 μL)
   2. 25 μL 3 M NaOAc
   3. 550 μL EtOH
   4. Incubate at -20ºC ≥ 1 hr.
4. Spin 5' to pellet DNA. Wash with 70% EtOH 2x to remove salts, air dry.
5. Resuspend DNA pellet in 55 μL of Hyb buffer.
6. Melt probe by incubating at 72ºC x15 min.
7. Incubate at 37ºC x30 min. to allow CoT DNA to anneal to probe.
8. Place probe on ice until ready to use.

Meanwhile in Coplin jars:

1. Hydrate: 2x SSC x2-5 min.
   (Optional, to reduce cytoplasm bkgd: 10 mg/mL RNAse (DNAse-free) in 2x SSC, 37ºC x30 min.)
2. Denature in 2x SSC + Formamide (50% vol), 72ºC x2-5 min.
3. Dehydrate: Ice cold 70% EtOH, 85% EtOH, 95% EtOH, 100% EtOH, each 5 min.
   (Denaturing solution and EtOH washes can be reused if kept in bottles).
4. Dry off back of slide with Kimwipe, and air dry by propping on one end.
5. Add 10 μL of probe to a region of slide with a spot of cells and cover with a 18 mm glass slip.
6. Seal edges with rubber cement using a 5 mL syringe. (Optional warm cement at 37ºC to reduce viscosity)
7. Place slides in a humid chamber o.n. at 37ºC.

Day 4: Washes and probe detection.

1. Place 6 Coplin jars in 42ºC bath: 3 jars with 2x SSC + Formamide (50% vol), and 3 jars with 2x SSC only.
2. (Reagents can be reused if kept in screw capped bottles - note order of reagents as first buffer will be the most "dirty")
3. Warm blocking agent to room temperature (4x SSC, 5% non-fat dry milk, 0.02% Na Azide).
4. Remove slides from humid chamber. Remove rubber cement. Gently slide off glass slips (dip in PBS if necessary).
5. Wash slides:
   • 2x SSC + Formamide (50% vol). 42ºC, x3.
   • 2x SSC x5 min. 42ºC, x3.
6. Blocking step:
   • Wipe off back of slide.
   • Add 100 μL block solution (4x SSC, 5% milk, 0.1% Tween-20) to the slide in 2 spots. (To store blocking reagent add 0.025% NaAzide and refrigerate at 4ºC)
   • Cover with a large glass slip for 5'.
   • Gently slide off slip.
7. Labeling step:
   • Add 100 μL of antibody (e.g. 1/400 avidin-FITC, Vector labs) diluted in blocking solution,
   • Cover with large slip. Incubate r.t. x 30 min.
8. Wash step:
   • Gently slide off cover slip.
   • Wash in 2x SSC + 0.005% CHAPS at room temp. for 5 min. (Repeat 3x)
9. View on fluorescent microscope for signal. BAC probes probably do not need further signal amplification.
10. If a second probe is being used on the same specimen then repeat the Block - Labeling - Wash steps for each (e.g. with anti-digioxigenin Texas Red).
11. If a small probe was used or if the signal is too dim then the signal can be amplified further with additional rounds of:
    • Biotinylated goat anti-streptavidin antibody (Vector) 0.5 mg/mL. Dilute 1/100 in
    • Streptavidin-FITC.

(M. Fero, courtesy of the Porter lab)
Revised 2/07
Also see Wei-Ming and Emily's protocol  This reference link is broken.

Reagents
Stains (change after every 3rd use)
Harris hematoxylin: 1X stock. Anatech Lmtd. Cat #842. (616) 964-6450
Gill's Hematoxylin: See recipe.
Eosin: 1x stock. Anatech Cat #837.

Solutions (make fresh each day)
Acid alcohol: 75% EtOH + 1/2500 v/v concentrated HCl (156 mL 95% EtOH + 44 mL H2O + 80 µL HCl = 200 mL)
Ammonia Solution: 0.084% (w/v) NH4OH (200 mL H2O + 0.6 mL 28% w/v NH4OH stock).

Organics
Xylene x2 (Make at least 1 fresh)
100% EtOH (Make at least 1 fresh)
95% EtOH x2 (make both fresh)
80% EtOH x1 (make fresh = 168 mL 95% EtOH + 32 mL H2O)

Deparafinization

1. Xylene 3' (x2)
2. 100% EtOH (x2)
3. 95% EtOH (x1)
4. 80% EtOH (x1)
5. H2O (x2)
6. Replace the dirty xylene #1, above with fresh xylene.
7. Replace the dirty 100% EtOH #1, above with fresh 100% EtOH.

Staining
(Note: Unless otherwise specified, slides should be dipped 10 times in each solution)

1. Hematoxylin, 2 min (x1)
2. Running H2O x 2 min
3. Acid Alcohol x1 ("Differentiation" lightens the staining, especially outside the nucleus) Skip this differentiation step for progressive staining with the weaker Mayer's Hematoxylin stain.
4. H2O x1
5. Ammonia solution 10 dips (x1) (changes the stain from purple to blue)
6. Running H2O 5 min
7. 80% EtOH x1
8. Eosin 15"
9. 95% EtOH x2
10. 100% EtOH
11. Xylene 3 min x2

Mounting

1. Wipe off the xylene off the back of a slide on a paper towel
2. Using a clean glass rod add a drop of permount to the slide
3. Cover with a glass slip (use 2 drops of permount for large slips)
4. Tilt the slide on edge on a paper towel to remove extra xylene or permount
5. If bubbles are present gently squeeze them out by pressing on slip with a pencil eraser
6. Place slides on a paper towel to cure overnight

M. Fero — 3/13/98

Materials

L-lysine coated glass cover slips or charged glass slides
Neutral Buffered Formalin (Sigma HT50-128)
0.5% NP40 in PBS
(2.5 N HCl or 0.07 N NaOH for BrDU staining only)
Primary and secondary antibodies
Vectashield (Vector Labs)

Protocol

1. Grow cells on L-lysine coated glass slips or cytospin cells onto charged glass slides.
2. Fix cells for 5 min. in neutral buffered formalin.
3. Permeabilize the nucleus by incubating in 0.5% NP40 in PBS at r.t.
4. Rinse in 3 changes of PBS for a total of 10 minutes.
5. For BrDU staining denature the DNA by one of the following:
   1. soak in 2.5N HCl at 37°C for 15 min, or
   2. 0.07N NaOH for 2 min at room temp
6. Add 100 µL primary antibody (titer determined emperically ~10x the concentration used in a western). Cover with a glass slip and place in a humidified chamber at r.t. for 1 hr.
7. Float the coverslip off by dipping into a jar of PBS, and rinse as in 4).
8. Add secondary antibody as in 5) and 6) above. (e.g. FITC-conj goat anti-rabbit (1:1000) or Biotinylated isotype specific anti mouse for immunoperoxidase staining). Wash as before.
9. To minimize quenching of flourochrome mount with Vectashield (Vector labs) and cover with a glass slip.

(courtesy of Farr lab, UWMC)

Materials

Cold Acetone (-20ºC) in Coplin jars
PBS, BenchKote paper, plastic lids
Anti-digoxigenin-HRP diluent: 1% BSA in PBS (store 10 mL aliquots of BSA at -20ºC)
Anti-rat-digoxigenin diluent: 5% Milk powder, 10% mouse serum, 10% goat serum, 1% BSA in PBS
Primary and secondary antibodies (this had a link that is broken)
DAB: 10 mL DAB (3.8% w/v), 240 mL H2O, 13 mL Tris (1M pH 7.6), 60 µL 30% H2O2.
Alternate DAB: 10 mg DAB (Sigma D5905, 10 mg/Tab, $156) 60 mL PBS, 140 µL 3% H2O2.

Cutting Frozen Sections

1. Place specimens at -20ºC x20 min.
2. Squeeze a dolop of OCT onto a mounting block and place frozen specimen on block.
3. Let OCT harden at -20ºC x15 min.
4. Set knife angle to 7.5º.
5. Advancing block: Rotate wheel to bring knife forward with brake on.
6. Bring block up with wheel at back
7. Advance with rotator wheel
8. Rotate block so tissue is perpendicular to knife
9. Cut OCT block with razor to form a rectangle or a trapezoid narrower on top
10. Set anti-roll device - back off it bounces by knob at bottom

Staining Procedure

1. Incubate slides in cold (-20ºC) acetone x 20'.
2. Tap acetone off each slide and incubate in PBS at r.t. x5'.
3. Saturate BenchKote paper with PBS. Remove a slide from PBS and wipe edges of each slide with KimWipe to create a static barrier. Add 175 µL of 1º antibody to each slide. Cover with a lid x1hr.
4. Rinse by dipping in 2 staing jars containing PBS. Soak in a 3rd PBS jar x5 min.
5. Add 175 µL of 2º antibody at r.t. as before x5 min.
6. Rinse with PBS as before.
7. Add DAB solution at r.t. x5 min.
8. Rinse in PBS 2x.
9. Step slides through graded EtOH to dehydrate: 70%, 95%, 100% (x3). Toluene (or xylene) (x2).
10. Add Permount and coverslip.

Modified from C.H. Lin 5/2005

Materials

Anti-digoxigenin-AP conjugate (Roche #11093274910, 150U/200 uL, $168)
Levamisole (Sigma #T1512, 2 g, $10) (blocks endogenous phosphatase activity)
BCIP (Roche #11383221001, 3 mL, $49)
NBT (Roche #11383213001, 3 mL, $48)
Atlternate BCIP/NBT (Roche, #11697471001, 20 Tabs, makes 10 mL/Tab, $67)
Blocking Reagent (Roche #11096176001, 50 g, $104, or 10x Block Buffer #11585762001, 1 set, $147)
Alternate blocking reagents: Nonfat dry milk, mouse serum, goat serum.
Tris-HCl, NaCl, MgCl2

Solutions

Buffer 1: 100 mM Tris-HCl pH 7.6, 150 mM NaCl, store at room temp.
Blocking Buffer: 10% w/v Blocking Reagent in Buffer 1.
Alternate Blocking Buffer: 1% w/v nonfat dry milk, 1% mouse serum, 1% goat serum, 1% BSA, in Buffer 1.
Buffer 2: 100 mM Tris-HCl pH 9.5, 50 mM MgCl2, 100 mM NaCl (use pH 10 for blue color)
Buffer 3: Buffer 2 with 0.5 mg/mL levamisole (make fresh)
Color Detection Solution: 0.45 uL/mL NBT, 3.5 uL/mL BCIP, in Buffer 3. (Make fresh)
Alternate Color Detection Solution: 1 Tablet of BCIP/NBT, 10 mL Buffer 3. (Make fresh)
TE: 10 mM Tris base pH=8.0, 1 mM EDTA. Store at room temp.

Procedure

1. Cut frozen sections. Fix in -20ºC acetone briefly and air dry. Altertatively cut paraffin sections and deparaffinize with xylene and graded EtOH into PBS.
2. Block slides with 1/20x serum/PBS (use the appropriate blocking serum, i.e. same as 1º Ab), 10'. Wash in PBS 2' x3.
3. Add digoxigenin labeled 1º antiobody. Ususally 1:1000 (v/v) in PBS. Wash in PBS 2' x3.
4. Equilibrate slides in Buffer 1, RT x 5'.
5. Incubate slides in Blocking Buffer, RT x 30'
6. Incubate with anti-Dig-AP conjugate (1:2000 in Blocking buffer.), 2 hrs. RT, or o.n. at 4ºC.
7. Wash in Buffer 1, 10' (x3).
8. Inactivate endogenous peroxidase by incubating in Buffer3 for ≥ 5'.
   Incubate in Color Detection Solution for 20 - 60', until desired intensity is achieved. Protect from light.
   Stop color development with TE pH=8.0.
9. Post-fix with 4% paraformaldehyde for 20'.
10. Wash 3x in PBS.
11. Dehydrate with graded EtOH and xylene and then coverslip with Permount.

Fixation

The usual fixative for paraffin embedded tissues is neutral buffered formalin (NBF). 10% NBF contains 4% formaldehyde by weight and is therefore equivalent to 4% paraformaldehyde. NBF contains a pH buffer plus a preservative (methanol) which prevents the conversion of formaldehyde to formic acid. Because of the preservative, NBF has a shelf life of months, whereas 4% PF must be made fresh.

Optimal formalin fixation requires adequate time to form -CH2- crosslinks, generally ≥ 48 hrs, at room temperature, for 2 mm-thin sliced tissues. Inadequately fixed tissues will become dehydrated during tissue processing, resulting in hard and brittle specimens. Alcohol based fixatives generaly do not give good morphology but may be useful for special cases (such as BrdU staining). A particular challenge is immunostaining fixed specimens. In many cases formaldehyde fixation will prevent recognition of epitopes by the primary antibody. Occasionally, "antigen retrieval" procedures will improve results. Usually frozen sections are a better bet for immunostaining, but at the expense of morphology. An alternative approach, suitable for thin or porous tissues, is to perform immunohistochemistry on fresh tissues and then post-fix and embed the tissues in paraffin.

B5 is a formalin fixative that also contains 6% (w/v) mercuric chloride. It is gives superior morphology for lymphoid tissues, but care must be taken not to over-fix tissues (fixation time = 2-12 hours). Becuase it contains mercury, B5 should not be used routinely. Care must be taken to minimize volumes and collect all used solutions for hazardous waste disposal.

Decalcification (only necessary for samples containing bone):
After fixation, bone must be decalcified or else it won't cut on the microtome.

• After fixation, bone must be decalcified or else it won't cut on the microtome. Immerse tissue cassette in 11% formic acid with a stir bar overnight in a fume hood.
• Rinse in running water for 30- 60 minutes (the smell should be gone).

Storage in 70% Ethanol

After adequate fixation, tissues may be transferred to 70% ethanol and stored at 4°C for 1 week. This should only be done for a short time, prior to paraffin processing. The main goal is to avoid contamination of processor solutions with fixative.

Paraffin processing
The Shandon Hypercenter XP tissue processor takes the cassettes through a series of graded EtOH baths to dehydrate the tissues and then into xylene. Hot paraffin can then permeate the tissues:

• 70% Ethanol 20 min (x1)
• 95% Ethanol 20 min (x2)
• 100% Ethanol 20 min (x2)
• Xylene 20 min (x2)
• Paraffin (65°C) 30 min. (x1)
• Paraffin (65°C) 30 min (x1) with vacuum (applied manually on our machine).

If the processor is to be run overnight it should be programmed to hold on the first ethanol bath and not finish until the next morning so the specimens do not sit in hot paraffin longer than the time indicated. If specimens are fresh they may incubate in formalin in the first stage on the machine. It is important to not keep the tissues in hot paraffin too long or else they become hard and brittle. The vacuum helps to speed up the permeation of tissues by paraffin and helps get rid of any small air bubbles. Processed tissues can be stored in the cassettes at room temperature indefinitely.

Embedding tissues in paraffin blocks
Tissues processed into paraffin are melted by placing the entire cassette in 65°C paraffin bath for 15 minutes. Turn the heat block on to melt the paraffin one hour before adding the tissue cassettes. Also warm metal block molds on the hot plate. The paraffinzed specimens may be dissected with a razor blade and placed with the cut surface down towards the bottom of the mold. Hot paraffin is added to the mold and from the paraffin pot. Use heated forceps to orient the tissues in the mold. When the tissue is in the desired orientation add the labeled tissue cassette on top of the mold as a backing. Be sure there is enough paraffin to cover the face of the plastic cassette. Slide the mold off of the hot plate onto a cold aluminum heat sink. When the wax is completely cooled and hardened (~20 min.) the paraffin block can be popped out of the mold. If the wax cracks or the tissues are not aligned well, simply melt them again and start over. Tissue blocks can be stored at room temperature for years.

Sectioning tissues
Turn on the water bath and check that the temp is 35-37ºC. Use fresh deionized water (DEPC treated water must be used if in situ hybridization will be performed on the sections). Blocks to be sectioned are placed face down on an ice block or heat sink for 10 minutes. Place a fresh blade on the microtome. Insert the block into the microtome chuck so the wax block faces the blade and is aligned in the vertical plane. Set the dial to cut 4-10 µM sections. The blade should angled 4-6º. Face the block by cutting it down to the desired tissue plane and discard the paraffin ribbon. If the block is ribboning well then cut another four sections and pick them up with forceps or a fine paint brush and float them on the surface of the 37ºC water bath. Float the sections onto the surface of clean glass slides. If the block is not ribboning well then place it back on the ice block to cool off firm up the wax. If the specimens fragment when placed on the water bath then it may be too hot.
Place the slides with paraffin sections in a 65°C oven for 20 minutes (so the wax just starts to melt) to bond the tissue to the glass. Slides can be stored overnight at room temperature.

Deparafinization

The paraffin is removed from the section with xylene. If the tissues are to be stained with an aqueuous solution then the slides must rehydrated in graded ethanol baths. Unless a time is indicated the approach is to gently agitate the slides by repeated immersion ~20x in each bath:

Procedure

1. Xylene for 2 min. (x3 changes)
2. 100% EtOH (x2)
3. 95% EtOH (x1)
4. 80% EtOH (x1)
5. H2O (x1)

Hematoxylin and Eosin (H&E)

This is the most conventional stain for formalin fixed paraffin sections. The hematoxylin stains negatively charged nucleic acids (nuclei and ribosomes) blue. The eosin stains proteins pink. The hematoxylin or the eosin can also be used by themselves in more dilute form as counterstains for immunoperoxidase staining. To do this dilute the stain 1:4 with H2O or EtOH, respectively. Slides to be stained in must be washed in ethanol first as listed below for the conventional protocol:

Reagents

1. Harris hematoxylin: 1X stock. Anatech Lmtd.(616) 964-6450
2. Eosin: 1X stock. Cat #837.
3. Acid alcohol: 76.6% EtOH, 1/300 v/v conc. HCl. (230 mL EtOH, 70 mL H2O, 1 mL HCl)
4. Ammonia Solution: 0.084 w/v NH4OH (1 L H2O 3 mL 28% w/v NH4OH stock).

Procedure

1. Hematoxylin, 2 minutes (x1)<>
2. Running water (x1)
3. Acid alcohol (x1)
4. H2O (x1)
5. Ammonia solution (x1)
6. Running water 5 minutes (x1)
7. 80% EtOH (x1)
8. Eosin 15 seconds
9. 95% EtOH (x2)
10. 100% EtOH (x2)

Wright Giemsa

This is the conventional stain for blood smears and bone marrow cytology. It is usually performed on an automated slide stainer (see pathology or Bernstein lab).

Methyl Green

(2% (w/v) methyl green in 0.1 M NaOAc, pH 4.2)

Methyl green is a nuclear counter stain which works nicely for immunoperoxidase stained slides. It is difficult to control the intensity of the stain however since it washes out in both aqueous and organic solutions and this will depend on how quickly you mount the slides. Mix 918 mL of 0.1N acetic acid with 331 mL of 0.1 M NaOAc and adjust pH to 4.2 with NaOH. Add 25 gm of methyl green dye. Filter through Whatman #2 filter paper.

Procedure

1. H2O x 10-15 sec.
2. Methyl green x 5 min.
3. H2O (x2).
4. Air dry.

New Methylene Blue

This stain is useful for distinguishing reticulocytes from mature RBCs in the peripheral blood. Mix whole blood 1:1 (v:v) with New Methlylene Blue (Ricca Chemical Co., Arlington Heights, IL). Incubate 10 minutes. Count on hemocytometer.

Benzidine Stain

This is a specialized stain which identifies erythroid cells (RBCs and their precursors). It gives them a golden brown color.

Procedure

1. Methanol, 10-15 seconds.
2. Benzidine, 5 min: [1% w/v of 3,3' dimethoxybenzidine in methanol] (This is toxic stuff).
3. Peroxide 2.5 min. (1 vol 30% H2O2 plus 11 vol. 70% EtOH)
4. Dionized water wash, 2.5 min.
5. Hematoxylin stain, 1.5 min.
6. Rinse in tap water, 8 min.
7. Air dry.

Acetylcholine esterase (AChE) Stain

Procedure

1. Fix in 5% glutaraldehyde x 15 min.
2. H2O (x3)
3. Flood slide with fresh AChE stain. (5 mM NaOAc, pH5; 1mM glycine; 0.2 mM CuSO4; 1.15 mg/ml acetylthiocholine iodide: Sigma A5751)
4. Incubate in petri dish o.n.
5. Rinse in H2O (x3), air dry.

M. Fero — 3/24/04 — adapted from P. Soriano

Materials

Fixative
2% Paraformaldehyde (in PBS)

Staining solution
5 mM potassium ferricyanide,
5 mM potassium ferrocyanide,
2 mM MgCI,
0.01% sodium deoxycholate
0.02% Nonidet P-40 (NP-40)
in PBS.

X-gal Stock
40 mg/mL X-Gal (5 -Bromo-4-chloro-3-indolyl-beta-D-galactopyranoside)
in dimethylformamide
store at -20°C in small aliquots.

X-gal Mix (make fresh just before use)
10 mL Staining solution
0.25 mL X-gal Stock (final = 1 mg/mL).
Be sure that X-Gal is well dissolved before use otherwise crystals may form in the staining solution

Procedure

1. Fix cells on ice for the following amount of times:
   15': Cells, small organs (pituitary, adrenal, thyroid), E8 embryos.
   30': Organ slices (2-3 mm), E10 embryos.
   60': Organ slices (4 mm), E12 embryos.
   90': Whole organs, E14 embryos.
2. Cut organs on a Vibratome in ice cold PBS to a thickness of 0.5 mm.
3. Transfer tissues to X-gal mix in small dishes or multi-well plates. Incubate at 37°C with gentle agitation until desired level of staining is achieved. Stain longer if histologic sections will be cut.
4. Place tissue slices on a small square of lens paper and transfer to histology tissue cassettes. The lens paper helps to keep the specimen from curling up during paraffin processing.

For your convenience, you can download the manual from the Bio-Rad website.

For your convenience, you can download the Direct PCR Tail DNA Preps manual from Vaigen Biotech.

For your convenience, you can download the manual here:

ImageJ-guide

For your convenience, you can download the manual here:

RNeasy_Mini_Handbook

For your convenience, you can download the manual here:

HB6_rotor_specs

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TRIzol_Reagent

Download the formulae here:

Formulae Document

A common issue in designing a clinical or animal study is to determine the number of subjects required. Power calculations provide an estimate of the number of study subjects that will be necessary to be able detect a difference between experimental and control subjects. The numbers required will depend on the magnitude of the difference between the two groups. For some experiments the primary measurement is the proportion of individuals with a certain outcome (e.g. yes/no or live vs. dead). The following graph depicts the results of power calculations using Fisher's exact method which is better for small numbers than a chi-square test. The data was generated with G*Power3 software. A different type of power calculation should be done if a numerical outcome is being measured for each individual (e.g. survival time) not simply a proportion.

The variables mentioned in the graphs are:

• N1 + N2 (the number of subjects in the experimental and control groups, combined)
• P1 (the proportion of experimental subjects which are positive for the endpoint)
• P2 (the proportion of control subjects which are positive for the endpoint)
• alpha ≤ 0.05 (the p-value of the study, the probability that such results might occur by chance)
• power = 0.8 (1- ß) (80% chance of being able to detect a difference between the groups)

Example: Suppose that you have strain of lab mice in which 10% develop tumors by 1 year of age (P2 = 0.1, see curve with red triangles). If your experimental group develops a higher frequency of tumors, say 40% (P1 = 0.4) then the total number of animals needed in the study (N1 + N2, on the Y-axis) is 60, or about 30 in each group. Notice how the curves rise sharply as the proportion of positive animals in the experimental group (P1) decreases.

1. Determine the final volume (Vf): How much to you want to make?
2. Determine the dilution factor (d.f.). User either...
   d.f. = Cs / Cf i.e. the ratio of starting concentration / final concentration:
   d.f. = the fold concentration (e.g. 20x) of the starting material (if the final concentration is 1x)
3. Calculate the starting volume (Vs): How much of the concentrated stock will you use?
   Vs = Vf / d.f.
4. Calculate the diluent volume (Vd): With how much water will you dilute the stock?
   Vd = Vf - Vs

Example #1: You want to make up 1 mL of oligonucleotide N1 at a concentration of 2 µM using a stock of 50 µM using water as the diluent.

1. Vf = 1000 µL
2. d.f. = Cs / Cf = 50 / 2 = 25x
3. Vs = Vf / d.f. = 1000 / 25 = 40 µL
4. Vd = Vf - Vs = 1000 - 40 = 960 µL
   1. Thus add 40 µL of the N1 primer + 960 µL H2O.

Example #2: You want to make up 7 L of TAE buffer from a 20x stock using water as the diluent.

1. Vf = 7000 mL
2. d.f. = 20x
3. Vs = Vf / d.f. = 7000 / 20 = 350 mL
4. Vd = Vf - Vs = 7000 - 350 = 6650 mL
   1. Thus use 350 mL of 20x TAE + 6650 mL H2O.