Protocols

Table of Contents

Cloning your Transgenic Vector                                                                                       

Preparing DNA vector                                                                                                    

Transformation of competent bacteria                                                                           

Cloning your vector                                                                                                        

Designing oligonucleotide linkers                                                                                   

Annealing linkers                                                                                                            

Ligation of linkers to cDNA                                                                                             

Restriction analysis of your vector                                                                                

Sequencing your vector                                                                                                

Generating transgenic or targeted ES cells                                                                     

Electroporation                                                                                                               

Drug selection of ES cells                                                                                              

Picking colonies                                                                                                              

Replica plating 96-well plates                                                                                        

Freezing 96-well plates of colonies                                                                               

Thawing cells from 96-well plates                                                                                 

LacZ staining of ES cells                                                                                               

Human alkaline phosphatase (hPLAP) staining of ES cells                                            

eGFP visualization of ES cells                                                                                        

ES cell DNA extraction and purification from 96-well plates                                         

ES cell DNA extraction and purification from 24-well plates                                         

Southern hybridization                                                                                                   

Cre excision to test cDNA and second reporter expression                                         

Genotyping mice                                                                                                               

LacZ staining                                                                                                                 

Human alkaline phosphatase (hPLAP) staining                                                              

eGFP                                                                                                                              

Staining tissue sections                                                                                                    

LacZ tissue staining                                                                                                       

Human alkaline phosphatase (hPLAP) staining                                                              

eGFP                                                                                                                              

Sorting eGFP positive cells                                                                                              

 

Cloning your Transgenic Vector

Preparing DNA vector

Your DNA vector is shipped on Whatman paper. To derive the DNA:

• Cut out the circular region containing the spotted DNA.
• Place inside a 1.5 mL microfuge tube.
• Add enough TE buffer to dampen the Whatman paper (approximately 100 mL; incubate at room temperature 15 min. The DNA will dissolve in TE.
• Collect TE with DNA by centrifugation: 14 000 rpm, 1 min.
• DNA is ready for transformation in competent bacteria.

Transformation of competent bacteria

• Thaw competent cells (eg. DH5a) in polypropylene tubes, on ice.
• Add DNA in TE (maximum 1/10 volume of cells) to cells.
• Incubate on ice 30 min.
• Heat shock cells at 42°C for 45 sec.
• Incubate on ice 2 min.
• Add SOC medium to cells and recover in shaking incubator at 37°C for 1 hour.
• Plate <200 mL on LB + ampicillin (100 mg/mL) selective plates. Incubate overnight at 37°C.
• Colonies can be grown in liquid culture for DNA extraction.

Cloning your vector

Multiple methods exist for subcloning a cDNA of interest into expression vectors. You may require oligonucleotide linkers to generate compatible restriction sites between the cDNA and the transgenic vector.

Designing oligonucleotide linkers

• Identify unique restriction sites flanking the cDNA.
• Design linkers that can anneal to the ends of the cDNA. The linker will contain on one end, the restriction site for the cDNA, and on the other, the restriction site for the cloning site of the transgenic vector.

G A A T T C C T C G A G G A A T T C

C T T A A G G A G C T C C T T A A G

        EcoRI            KpnI              EcoRI

 

For example, if an EcoRI site is at the 3’ end of the cDNA, and you want to subclone into the KpnI site of the transgenic vector, the linker would look like the red and blue portion of the above oligonucleotide. The black portion represents the 3’ end of the cDNA. Therefore you would order a 12-mer that is 5’-AATTCCTCGAGG-3’.

Note that in cloning a cDNA into iZAP or iZEG, you do not need to put the cDNA in-frame with the Alk Phos or EGFP, because the Alk Phos and EGFP are preceded by an IRES sequence.

Annealing linkers

• The linker is self-annealing in the following conditions.
• Dilute linker to 1 mg/mL with annealing buffer: 10 mM Tris-HCl pH7.5-8; 50 mM NaCl; 1 mM EDTA
• Heat to 94°C 3-5 min.
• Cool at room temperature. Store at -20°C.

Ligation of linkers to cDNA

• Digest 10 mg cDNA vector with restriction enzyme in 100 mL volume
• Extract DNA with chloroform and precipitate with ethanol.
• Resuspend DNA in £12 mL TE.
• Add 1 mL annealed linker, 4 mL water, 2 mL T4 ligase buffer, 1 mL T4 ligase.
• Incubate 12°C for 4 hours.
• Remove 1 mL of the ligation for analytical gel. Digest the remaining ligation with the restriction enzyme for the transgenic vector cloning site (in the example above, KpnI). Use an excess of restriction enzyme (100 U) in 100 mL volume to digest the excess linker. Digest overnight.
• Extract DNA with chloroform and precipitate with ethanol.
• Gel purify the cDNA. We recommend double gel electrophoresis of the cDNA. After the first run, excise the band containing your cDNA, and insert the gel slice into a new agarose gel. Gel purify the cDNA fragment from the second electrophoresis. The second electrphoresis step removes any remaining linker from the cDNA.
• The cDNA can be ligated to the transgenic vector.

Restriction analysis of your vector

Restriction enzyme digests may be used to confirm the insertion and orientation of the cDNA insert in the transgenic vector.

Sequencing your vector

The vector can be sequenced to confirm the position of your cDNA relative to the expression of the reporter gene.

Generating transgenic or targeted ES cells

The transgenic vector is ideally introduced into murine embryonic stem (ES) cells through electroporation or lipofectin transfection methods. We recommend using ES cells rather than pronuclear injection to generate transgenic mice, because you can select ES cell clones with strong lacZ expression and a single-copy transgene. This provides widespread expression in the mice you generate and avoids recombination between multiple tandem loxP sites. Remember to use feeder cells that are neomycin- or puromycin-resistant, depending on which drug you are selecting with.

Electroporation

Day 0: Prepare plates of freshly inactivated (mitomycin C-treated) feeders in PMEF medium.

Day 1: Thaw one vial of ES cells (an equivalent of one fifth of a 10 cm plate) onto two 10 cm plates with a confluent lawn of feeders in ES cell medium. Feeder plates can be used for several days after being prepared.

Day 2: Refeed ES cells (with cold medium). The cells should be well-defined colonies with clear edges.

Day 3: Passage cells 1:5 or 1:7 into 10 cm plates.

Day 4: Refeed ES cells. Prepare DNA for electroporation by linearizing the vector.

• Digest 10-20 mg DNA per cuvette with ~5 U restriction enzyme.
• Extract DNA using chloroform, then precipitate using ethanol. Resuspend DNA to a concentration of 1 mg/mL. Confirm the digestion and DNA digestion by electrophoresis of a small amount of DNA.

Day 5: Electroporate ES cells:

• Trypsinize cells at 37°C for 5 min or until the cells dissociate with gentle disruption of the plate.
• Collect the cells in 5 mL of medium, and use this volume to collect and pool the cells from the remaining plates into a 50 mL conical tube. Pellet the pooled cells by centrifugation at 1200 rpm for 5 min. Gently aspirate the supernatant and add one to two drops of PBS (-Ca²⁺/-Mg²⁺). Loosen the pellet by tapping/swirling the tube. Add 0.5 mL PBS, and count cells. Resuspend cells to 7 x 10⁶ cells/mL. A confluent plate of cells should give up to 10⁷ cells.
• In the cuvette, add 20 mg DNA (200 mL) and 0.8 mL cells. Electroporate. Immediately place cuvette on ice for 20 min to allow cells to recover.
• Gelatinize the number of 10 cm plates required, and add 10 mL media to each plate.
• Transfer electroporated cells into gelatinized plates. Each cuvette of 5.6 x 10⁶ cells can be split into 2-4 10 cm plates. Place in 37°C incubator.

Day 6: Refeed ES cells with normal medium. There should be ~50% cell death observed.

Drug selection of ES cells

Day 7: Drug selection with either G418 or puromycin should begin 48 hour after electroporation or transfection. Use the drug concentration pre-determined by a dose response curve.

Day 8: Refeed the cells with freshly added drug.

Day 9: Refeed. There should be obvious cell death. Drug-sensitive colonies have edges that become less sharp, and cells that are dead on top. Drug-resistant colonies have clear edges and look shiny.

Day 11: There should be massive cell death. Refeed with medium containing drug.

Day 12: There will be some colonies that continue to die. Resistant colonies should be more visible as round, nut-shaped colonies. The edges of colonies should be sharp, with no brownish centre. Colony sizes may vary.

Day 13: Some residual cell death may be observed. Two types of resistant colonies should be observed: fast growing colonies that are visible to the naked eye, and smaller colonies that grow 3-4 days slower than do the large ones. The fast growing colonies may have varying morphologies: tightly-packed colonies (OK), dense centred but with a peripheral rim of flattened cells (OK) which may be fairly large (OK), or completely flattened colonies (not suitable). The slower-growing colonies are likely resistant, but express less drug resistance genes. These colonies are not typically used for injection.

Day 14: Refeed cells.

Day 15: Refeed cells.

Picking colonies

To screen individual ES cell colonies for reporter expression, pick single colonies into individual wells of 96-well plates. By day 16, colonies should be large enough to be picked.

• Prepare the 96-well plates.  Two 96-well plates are needed for each colony you pick: one flat-bottom plate, and one V-bottom intermediate plate for cell trypsinization. Gelatinize the flat-bottom plate and add 100 mL ES cell media with drug in each well. To the V-bottom plate, add 40 mL trypsin.
• Replace the media in the 10 cm plates of colonies with 5 mL PBS. Pick colonies from the 10 cm plates by using a dissecting microscope in the laminar flow hood.
• To pick a colony, use a P20 set to 15 mL. Using the tip, scrape a colony off of the plate while aspirating. The cells should transfer from the plate to the pipet tip.
• Transfer the cells from the tip to one well of trypsin. Pipette the cells 2-3 times to ensure all cells have been transferred.
• Leave cells in trypsin 20-40 min, then add 60 mL ES medium to the 40 mL of trypsinized cells. Transfer this 100 mL to the 100 mL media in the flat-bottom 96-well plate. A multi-channel pipette will be useful for this step. Incubate the cells at 37°C.

Day 17: Refeed the colonies early the next day.

Day 18: Refeed the picked colonies with fresh ES cell medium with drug.

Day 19: Refeed the picked colonies with fresh ES medium with drug.

Replica plating 96-well plates

Day 20: When ES cells in the 96-well plate become confluent, passage them 1:3.

• Aliquot 100 mL ES medium into wells of three 96-well plates. One plate should contain feeder cells, which will be frozen; two plates should be gelatinized. One plate can be used for checking reporter expression of DNA extraction, and one plate can be used for propagation.
• Wash confluent ES cells with PBS (Ca-/Mg-), and trypsinize by adding 50 mL trypsin to each well. Incubate at 37°C for 5 min. Add 100 mL ES cell medium to each well of cells, then aliquot this 150 mL evenly to the three new plates.

Day 21: Feed cells with fresh medium.

Day 22: Refeed the cells. Place a Styrofoam box at -80°C in preparation for freezing the ES cells on feeders.

Freezing 96-well plates of colonies

Day 23: Two hours prior to freezing, change the media.

• Prepare 2X freezing media: 30% FCS, 20% DMSO in DMEM. Place on ice.
• Wash cells once with 200 mL PBS.
• Add 50 mL trypsin to each well. Incubate at 37°C for 5-10 min.
• Place plate on ice, and add 50 mL ES cell medium.  Pipette up and down ten times.
• Add 100 mL 2X freezing medium.
• With the plate still on ice, add 80 mL cold sterile mineral oil. Wrap plate with parafilm, then with foil. Place plate in the Styrofoam box at -80°C.

Thawing cells from 96-well plates

It is essential to rapidly thaw cells and get them into fresh ES medium.

• Prepare 24-well plates with feeders.
• Remove the plate from -80°C and warm the plate in your hands or on the surface of a 37°C water bath.
• Aspirate the mineral oil from each well.
• Pipette the cells up and down 2-3 times to resuspend the cells.
• Transfer to 24-well plate and incubate at 37°C.

LacZ staining of ES cells

Cells can be stained for lacZ expression one day after passage. The positive clones (typically 10% of total) can be passaged into two 24-well plates. One 24-well plate (gelatinized) can be used for DNA extraction, and the other 24-well plate (with feeders) is passaged into 6-well plates, then 10 cm plates. These cells are for frozen stocks.

• Wash cells once with room temperature PBS.
• Fix cells with 4% paraformaldehyde at room temperature for 5-10 min.
• Wash cells three times with room temperature PBS.
• Add 50-100 mL staining solution to each well. Incubate 4 hours to overnight at 37°C in the dark.
• Replace staining solution with equal volume of PBS.
• Store at 4°C if needed to intensify lacZ staining. LacZ-positive colonies are blue.

X-gal staining solution: store unused solution wrapped in foil at -20°C. IMPORTANT: Start with PBS first, or else the other components will not dissolve.

Final concentration	Stock concentration	To make 10 mL
1X PBS	1X	9.35 mL
1 mg/mL X-gal in DMSO	40 mg/mL	250 mL
5 mM K3Fe(CN)6	500 mM	100 mL
5 mM K4Fe(CN)6	500 mM	100 mL
2 mM MgCl2	100 mM	200 mL

Human alkaline phosphatase (hPLAP) staining of ES cells

• Wash cells once with PBS.
• Fix cells with 4% paraformaldehyde at room temperature for 5 min.
• Wash three times with PBS.
• Inactivate endogenous AP: securely wrap plate in parafilm and float on 65°C water bath for 30 min.
• Stain by adding NBT/BCIP (Sigma-Aldrich B1911) with 1X stain solution (see below). Incubate in the dark ½ - 4 hours at room temperature.
• AP positive colonies should be purple.

 

NBT/BCIP stain solution: make 20X in ddH₂O, then dilute 1:20 and add NBT/BCIP.

20X concentration

2 M NaCl

0.2% sodium deoxycholate

0.4% NP-40

eGFP visualization of ES cells

eGFP expression can be assayed in live cells with a fluorescent microscope. 

ES cell DNA extraction and purification from 96-well plates

DNA from this procedure is used for Southern hybridization to check for single copy integration of the transgene. See below for extraction from 24-well plate (which is more reliable to work).

• Remove ES medium with multi-channel pipette or aspirator.
• Wash once with 100 mL PBS.
• Add 50 mL cell lysis buffer (see below). Wrap in paraffin and incubate at 50°C overnight.
• Next day, add 100 mL cold absolute ethanol.
• Incubate at room temperature.
• Pellet DNA by centrifuging the plate at 10°C, 1000 rpm, 5 min.
• Invert the plate on paper towels to remove ethanol.
• Wash three times with 150 mL 70% ethanol. After the last wash, vigorously pat the plate on paper towels to remove as much ethanol as possible.
• To dissolve DNA, add 25 mL 0.1X TE. Seal the plate to prevent evaporation, and leave at room temperature overnight. Store plate at 4°C until you are ready to perform the Southern.
• The day before you start the Southern, digest the DNA overnight in a total volume of 30 mL.
• Run the samples overnight on a 0.8% agarose gel, 20-25V, and proceed with Southern hybridization.

Lysis buffer. Make fresh before use.

Final concentration	Stock concentration	To make 10 mL
0.5% sarcosyl	10%	500 mL
200 mM NaCl	5 M	400 mL
10 mM EDTA (pH 8.0)	500 mM	200 mL
10 mM Tris-HCl (pH 8.0)	1 M	100 mL
1 mg/mL proteinase K	20 mg/mL	500 mL

ES cell DNA extraction and purification from 24-well plates

• Remove ES medium with aspirator.
• Wash once with 500 mL PBS.
• Add 500 mL lysis buffer (see below) per well and incubate at 37°C overnight.
• Next day, transfer the lysed cells to sterile 1.5 mL microfuge tubes containing 500 mL isopropanol.
• Mix by inverting for 1 min.
• Using a sterile pipette tip, scoop out DNA threads into 500 mL 70% ethanol. Pellet DNA by centrifugation.
• Dissolve DNA by adding 100-200 mL TE (pH 7.5). Incubate at 55°C for 1-2 hours or 4°C overnight.
• The day before you start the Southern, digest the DNA overnight in a total volume of 30 mL.
• Run the samples overnight on a 0.8% agarose gel, 20-25V, and proceed with Southern hybridization.

 

Lysis buffer. Make fresh before use.

Final concentration	Stock concentration	To make 10 mL
1% SDS	20%	500 mL
100 mM NaCl	5 M	200 mL
1 mM EDTA (pH 7.5)	500 mM	20 mL
10 mM Tris-HCl (pH 7.5)	1 M	100 mL
500 mg/mL proteinase K	20 mg/mL	250 mL

Southern hybridization

Day 1: Restriction enzyme digest

Digest 10 mg genomic DNA in a total volume of 40 mL. Digests should be performed overnight.

Day 2: Gel electrophoresis

Run your samples on a 0.8% agarose gel overnight, at 20-25V. Add the appropriate amount of loading dye.

Day 3: Southern transfer. Adapted from Manniatis, 9.44.

• Take a picture of the gel with a ruler along the side with the molecular weight marker.
• Place the gel in a tray and rinse briefly in ddH₂O. Pour off water.
• Add enough 0.2N HCl to cover the gel. Set on gentle agitation at room temperature until the DNA loading dye color changes from blue to yellow.
• Pour off the HCl solution and rinse with ddH₂O.
• Cover the gel with denaturation buffer. Shake with gentle agitation at room temperature 30-45 min or until the color of the DNA loading dye changes back to blue. Denaturation buffer: 1.5 M NaCl, 0.5 M NaOH.
• Rinse with ddH₂O.
• Cover the gel with neutralization buffer. Shake with gentle agitation at room temperature 30-45 min. Neutralization buffer: 1.5 NaCl, 0.5 M Tris-HCl pH 7.4.
• Pour off neutralization buffer, rinse with ddH2O and add fresh neutralization buffer. Shake with gentle agitation for 15 min.
• Set up the capillary blot. Use 10X SSC or 10X SSPE as blotting buffer. Use 3 MM Whatman paper as the wick; ensure it is wet and that any bubbles are removed by rolling the paper with a 5 mL pipette.
• Place the gel on the wick, with the DNA side up.
• Squeeze out any bubbles between the gel and the wick with a glass pipette.
• Surround the gel with plastic wrap to prevent evaporation of the blotting buffer and to direct the absorption of the blotting buffer by the paper towels.
• Cut a piece of Hybond N+ membrane the same size as the gel. Cut one of the corners to mark the orientation of the membrane relative to the gel.
• Cut four pieces of 3 MM Whatman paper that are the same size as the gel.
• Place the Hybond N+ on the gel. Squeeze out bubbles with a glass pipette.
• Wet one piece of Whatman paper in blotting buffer and place on top of the Hybond N+.
• Place the other three pieces (dry) on top of the wet Whatman paper.
• Cover the surface of the Whatman paper with a 5 cm tall stack of paper towels.
• Place a 0.75-1 kg weight on top. Allow transfer to proceed overnight.

Day 4: Prehybridization and hybridization

• Dismantle the transfer apparatus.
• Before removing the gel from the membrane, turn the membrane gel-side up so that the membrane is on the bottom and the gel is on top of it, upside down. Mark each well of the gel on the membrane by placing the pencil through the gel. The marked side is also the side with the DNA.
• Rinse membrane in 2X SSC or 2X SSPE, whichever was used as blotting buffer.
• Place membrane on plastic, with DNA side up.
• Allow the membrane to air dry, then place the membrane and plastic wrap (with DNA side up) in the UV cross-linker.
• Completely cover the membrane with plastic wrap, and place it in a folded sheet of Whatman paper and store at 4°C without bending.
• Prepare the pre-hybridization solution fresh, or thaw a frozen aliquot.
• Heat the required volume of pre-hybridization solution to 42°C.
• Meanwhile, place the membrane in 6X SSPE until it becomes thoroughly wet, at least 2 min.
• Slip the wet membrane into a glass tube. Squeeze out any air bubbles between the membrane and the glass with a glass pipette.
• Add the pre-hybridization solution. Close the tube.
• Incubate in a rotating oven at 42°C 1-2 hours.
• Prepare hybridization solution or thaw an aliquot and heat. The solution should be heated to 42°C solution if the solution contains formamide, and 65°C if it does not contain formamide.

Pre-hybridization solution. Store at -20°C. Use 0.2 mL solution/cm² of Hybond N+ membrane.

Final concentration	Stock concentration	To make 25 mL
5X SSPE	20X	6.25 mL
5X Denhardt’s	50X	2.50 mL
1% SDS	10%	2.50 mL
0.1 mg/mL Denatured salmon sperm DNA	1 mg/mL	2.50 mL
50% Formamide		11.25 mL

 

20X SSPE

175.3 g NaCl

27.6 g NaH₂PO₄-H₂O

7.4 g EDTA

Add 800 mL ddhH₂O

Adjust the pH to 7.4 with ~6.5 mL 10 N NaOH

Make up the volume to 1 L

Autoclave

Hybridization solution. Store at -20°C.

Final concentration	Stock concentration	To make 25 mL
5X SSPE	20X	6.25 mL
1% SDS	10%	2.50 mL
0.1 mg/mL Denatured salmon sperm DNA	1 mg/mL	2.50 mL
10% Dextran sulphate		2.50 mL
50% Formamide	100%	11.25 mL

 

Probe preparation

Prepare probe according to the instructions in your kit.

Probe purification

• Plug the bottom of a 1 mL disposable syringe with a small amount of glass wool.
• Using as Pasteur pipette, fill the syringe with Sephadex G-50 equilibrated in 1X TEN buffer pH 8.0.
• When the syringe is full, place it in a 15 mL snap-cap tube.
• Centrifuge at 1500 rpm, 5 min at room temperature.
• Discard the eluate. Apple the DNA sample to the syringe barrel in a total volume of 100 mL.  Place the syringe into a 15 ml snap-cap tube.
• Centrifuge at 1500 rpm, 5 min at room temperature. The probe will collect in the bottom of the snap-cap tube. There should be close to 100 mL recovered.
• Transfer the probe to a 1.5 mL microfuge tube. Measure the radioactivity of the probe by Cherenkov counting: place 1 mL of probe into a 1.5 mL microfuge tube, close it tightly, place into a scintillation vial, and count. The count should be ~10⁶ cpm.
• Denature the probe by heating at 95°C for 5 min, then place in ice for 5 min.

Hybridization

• Remove the tube with the membrane from the incubator.
• Discard the pre-hybridization solution.
• Add the pre-warmed hybridization solution. Squeeze out bubbles with a glass pipette.
• Add the probe, but avoid direct contact with the membrane. Swirl the tube. Tightly close the tube.
• Incubate in rotating oven at either 42°C(with formamide) or 65°C (no formamide) overnight.

Day 5: Wash

• Remove the tube from the rotating oven.
• Carefully decant the hybridization solution into the appropriate radioactive waste container.
• Remove the membrane and place in a clean plastic container. Wash with 2X SSPE, 1% SDS at room temperature for 10 min. Repeat.
• After the second wash, replace with 1X SSPE, 1% SDS. Incubate at 65°C for 15 min.
• Wash membrane with 0.1X SSPE, 1% SDS at 65°C for 15 min.
• Decant the wash solution and check radioactivity. Repeat last wash step if the membrane is too hot.
• Wrap the membrane in plastic wrap, making sure to smooth out any bubbles.
• In the dark room place the membrane, DNA side up, on the back of an autoradiography cassette, and place autoradiography film on top, followed by an intensifying screen.  Close the cassette and place at -70°C 1-3 days.
• Develop the film. Alternatively, a phosphoimager may be used to capture the signal.

Cre excision to test cDNA and second reporter expression

Cre expression in lacZ positive ES cell clones can be used to confirm excision of lacZ and expression of the cDNA and the second reporter gene. This protocol is optimized for ES cells, and adapted from the protocol for LIPOFECTAMINE 2000 (Invitrogen).

• One day before transfection, seed 2 x 10⁵ ES cells carrying a single copy of the transgenic vector (as determined by lacZ stain and southern blot) in a 6-well gelatinized plate.
• Prepare the following solutions for each transfection:
• 1 mg linearized Cre DNA per transfection, in 100 mL serum-free medium or OPTI-MEM I Reduced Serum Medium. (An example of Cre vector to use is pCX-NLSCre-PGK-puro)
• 10 mL LIPOFECTAMINE 2000 in 100 mL serum free medium or OPTI-MEM.
• Combine the two solutions, mix gently, and incubate at room temperature for 45 min to allow DNA-liposome complexes to form.
• Rinse ES cells once with 2 mL serum free DMEM medium (no FCS).
• Add 0.8 mL serum free medium to the DNA-liposome complexes. Mix gently and overlay onto the ES cells.
• Incubate for 5 hours at 37°C in a CO2 incubator, then add 1 mL ES medium with 30% FCS.
• Next day, change the media.
• Start drug selection for Cre expression 48 hours after transfection. Use a drug concentration appropriate for your cell line. For puromycin, we use 1.5 ug/ml puromycin.
• Change the medium and add fresh drug daily.
• Approximately 30-40 colonies will form in one 6-well transfection. At day 7 after selection, staining/visualization of either lacZ and alkaline phosphatase or eGFP can be performed.

Genotyping mice

Mice may be genotyped by Southern blot and probing for a fragment of the transgenic vector. Mouse earclips can be directly stained for b-galatosidase or alkaline phosphatase expression. eGFP can be directly visualized in earclips under a fluorescent microscope.

LacZ staining

• Place mouse earclips into individual wells of 96-well plate.
• Wash 3 times with 100 mL PBS.
• Fix 10 min at room temperature with 100 mL of 0.2% glutaraldehyde in PBS.
• Wash 3 times with 100 mL PBS. After the last wash, remove as much PBS as possible.
• Add stain solution (see above). Add 100 mL, or enough to completely submerge the earclip. Cover plate, and incubate at 37°C in the dark 15 min-2 hours. LacZ positive earclips will be blue.

Human alkaline phosphatase (hPLAP) staining

• Place mouse earclips into individual wells of 96-well plate.
• Fix 30’ on ice with 100 mL of 0.2% glutaraldehyde in PBS. For better penetration of the Alkaline Phosphatase substrates, 0.02% NP-40 and 0.01% sodium deoxycholate can be added to the fix solution. Shake plate periodically.
• Wash 3 times with 100 mL PBS.
• Preheat PBS to 70°C. Add 100 mL PBS to each earclip. Inactivate endogenous AP by incubating at 70°C for 30 min. The plate may be sealed and floated on top of a 70°C water bath.
• Rinse once with PBS.
• Wash 3 times for 5 min with 100 mL AP wash buffer.
• Add 100 mL NBT/BCIP (Sigma-Aldrich B1911) with 1X NBT/BCIP stain solution. Place plate in room temperature, away from light. Incubate 30 min- 2 hours. hAP positive earclips will be purple.

AP wash buffer

Stock concentration	To make 100 mL
1 M Tris-HCl pH 9.5	10 mL
5M NaCl	2 mL
1 M MgCl2	1 mL
ddH20	87 mL

 

NBT/BCIP stain solution: make 20X in ddH₂O, then dilute 1:20 in NBT/BCIP.

20X concentration

2 M NaCl

0.2% sodium deoxycholate

0.4% NP-40

 

eGFP

eGFP fluorescence can be visualized in earclips and live animals using an appropriate light source and filter.

 

Staining tissue sections

LacZ tissue staining

• Dissect samples into cold PBS.
• Fix samples: Add fix solution and shake on ice for 4 hours. After the first hour, bisect the tissue to allow the fix solution to penetrate.
• Wash 3 times for 15-20 min with lacZ wash buffer.
• Cryoprotect samples in 15% sucrose in PBS for 1 hour at 4°C.
• Replace 15% sucrose solution with 30% sucrose in PBS overnight at 4°C.
• Next day, place tissue in Tissue-Tek OCT (Sakura) at 4°C for at least 1 hour.
• Embed the tissue in OCT: Place OCT into a labeled plastic mold.
• Place the tissue into the mold, and orient it.
• Place the mold onto dry ice. Hold the mold level, being careful to maintain the orientation of the tissue.
• When all samples have been prepared, wrap them in plastic wrap and store at -80°C.
• Prior to sectioning, allow blocks to equilibrate to -20°C in the cryostat chamber.
• Cryosection the tissue blocks at 10 mM. Place the sections onto poly-lysine coated slides.
• Dry slides 1-4 hours at room temperature.
• Store at -20°C in a slide box, with a small amount of dessicant.
• To stain for lacZ, equilibrate the slides to room temperature before removing them from the box.
• Place the slides into a staining jar.
• Fix slides in lacZ fix solution for 10 min.
• Wash 3 times for 5 min with lacZ wash buffer.
• Stain slides in lacZ staining solution for 4-6 hours, 37°C, protected from light.
• Rinse slides 3 times for 5 min in PBS.
• Dehydrate slides:
• 5 min PBS
• 5 min 70% ethanol in PBS
• 5 min 90% ethanol in PBS
• 5’ 100% ethanol
• 5’ 1:1 100% ethanold:xylene (do this step in fume hood)
• 5’ xylene (do this step in fume hood)
• 5’ xylene (do this step in fume hood)
• Mount with coverslips. Use a seal such as Cytoseal.

LacZ fix solution

Stock concentration	To make 50 mL
25% glutaraldehyde	0.4 mL
250 mM EGTA pH7.3	1.0 mL
1 M MgCl2	5.0 mL
PBS	43.5 mL

 

LacZ wash buffer

Stock concentration	To make 500 mL
1 M MgCl2	1.0 mL
1% sodium deoxycholate	5.0 mL
2% NP-40	5.0 mL
PBS	489 mL

 

LacZ stain buffer

Stock concentration	To make 75 mL
LacZ wash buffer	72.0 mL
25 mg/mL X-gal (in DMSO)	3.0 mL
K3Fe(CN)6	0.159 g
K4Fe(CN)6	0.123 g

Human alkaline phosphatase (hPLAP) staining

• Dissect samples into cold PBS.
• Fix samples: Add fix solution and shake on ice for 4 hours. After the first hour, bisect the tissue to allow the fix solution to penetrate.
• Wash 3 times for 15-20 min with lacZ wash buffer.
• Cryoprotect samples in 15% sucrose in PBS for 1 hour at 4°C.
• Replace 15% sucrose solution with 30% sucrose in PBS overnight at 4°C.
• Next day, place tissue in Tissue-Tek OCT (Sakura) at 4°C for at least 1 hour.
• Embed the tissue in OCT: Place OCT into a labeled plastic mold.
• Place the tissue into the mold, and orient it.
• Place the mold onto dry ice. Hold the mold level, being careful to maintain the orientation of the tissue.
• When all samples have been prepared, wrap them in plastic wrap and store at -80°C.
• Prior to sectioning, allow blocks to equilibrate to -20°C in the cryostat chamber.
• Cryosection the tissue blocks at 10 mM. Place the sections onto poly-lysine coated slides.
• Dry slides 1-4 hours at room temperature.
• Store at -20°C in a slide box, with a small amount of dessicant.
• To stain for hAP, equilibrate the slides to room temperature before removing them from the box.
• Place the slides into a staining jar.
• Fix slides in lacZ fix solution + 0.02% NP-40 + 0.01% sodium deoxycholate for 10 min.
• Wash 3 times for 5 min with PBS.
• Inactivate endogenous AP by incubation in PBS at 70°C for 30 min in a preheated staining jar. Let slides cool to room temperature.
• Rinse once with PBS.
• Wash in AP buffer for 10 min.
• Shake excess liquid from slides and place in a plastic box, tissue side up.
• Apply 0.5-1 mL NBT/BCIP (Sigma-Aldrich B1911) with 1X NBT/BCIP stain solution (see below) to each tissue sample. Close the box, and cover with foil to protect from light.
• Incubate at room temperature 10-30 min.
• Wash 3 times for 10 min in PBS + 2 mM MgCl₂
• Dehydrate slides:
• 5 min PBS
• 5 min 70% ethanol in PBS
• 5 min 90% ethanol in PBS
• 5’ 100% ethanol
• 5’ 1:1 100% ethanol:xylene (do this step in fume hood)
• 5’ xylene (do this step in fume hood)
• 5’ xylene (do this step in fume hood)
• Mount with coverslips. Use a seal such as Cytoseal.

AP wash buffer

Stock concentration	To make 100 mL
1 M Tris-HCl pH 9.5	10 mL
5M NaCl	2 mL
1 M MgCl2	1 mL
ddH20	87 mL

 

NBT/BCIP stain solution: make 20X in ddH₂O, then dilute 1:20 in NBT/BCIP.

20X concentration

2 M NaCl

0.2% sodium deoxycholate

0.4% NP-40

 

eGFP

eGFP can be detected in frozen sections. We have used anti-GFP, rabbit IgG fraction, HRP conjugate (Invitrogen A10260) primary antibody, Peroxidase goat anti-rabbit IgG secondary antibody (Vector Laboratories PI-1000), VECTASTAIN ABC Kit for goat IgG (Vector Laboratories PK-4005), and DAB Peroxidase Substrate Kit (Vector Laboratories SK-4100).

• Dissect samples into cold PBS.
• Fix samples in 4% paraformaldehyde.
• Cryoprotect samples in 15% sucrose in PBS for 1 hour at 4°C.
• Replace 15% sucrose solution with 30% sucrose in PBS overnight at 4°C.
• Next day, place tissue in Tissue-Tek OCT (Sakura) at 4°C for at least 1 hour.
• Embed the tissue in OCT: Place OCT into a labeled plastic mold.
• Place the tissue into the mold, and orient it.
• Place the mold onto dry ice. Hold the mold level, being careful to maintain the orientation of the tissue.
• When all samples have been prepared, wrap them in plastic wrap and store at -80°C.
• Prior to sectioning, allow blocks to equilibrate to -20°C in the cryostat chamber.
• Cryosection the tissue blocks at 10 mM. Place the sections onto poly-lysine coated slides.
• Dry slides 1-4 hours at room temperature.
• Optional: fix slides in 50% acetone/50% methanol for 10 min at -20°C.
• Air dry slides.
• Wash in PBS 5 min.
• Incubate slides for 30 min in 3% H₂O₂ in methanol to block endogenous peroxidase activity.
• Wash 4 times for 5 min with PBS.
• Block with 10% serum from the same animal as your secondary antibody.
• Incubate the anti-GFP antibody (1:1000) in blocking buffer overnight, at 4°C in a humidified chamber.
• Wash 4 times for 5 min with PBS.
• Incubate the secondary antibody at room temperature for 30 min in a humidified chamber.
• Make ABC according to VECTASTAIN protocol.
• Wash slides 4 times for 5 min in PBS.
• Incubate in ABC for 30 min in humidified chamber.
• Wash slides 4 times for 2-3 min in PBS.
• Make DAB solution according to DAB Peroxidase Substrate Kit protocol. Add to slides immediately. Color change should occur in 1 min.
• Drain slides and submerse in ddH₂O for 5 min.
• Counterstain with hematoxylin. Wash with running tap water for 4 min.
• Dehydrate slides:
• 5 min 70% ethanol in PBS
• 5 min 90% ethanol in PBS
• 5’ 100% ethanol
• 5’ 100% ethanol
• 5’ 1:1 100% ethanol:xylene (do this step in fume hood)
• 5’ xylene (do this step in fume hood)
• 5’ xylene (do this step in fume hood)
• Mount coverslips with Cytoseal and allow to dry overnight.

Sorting eGFP positive cells

Cells expressing eGFP may be sorted and collected using flow cytometry.