Generation and Validation of Tissue-Specific Knockout Strains for Toxicology Research

Abstract

Tissue-specific knockout mice are widely used throughout scientific research. A principal method for generating tissue-specific knockout mice is the Cre-loxP system. Here, we give a detailed description of the steps required to generate and validate tissue-specific knockout mice using the Cre-loxP system. Basic Protocol 1 describes how to use gene targeting in mouse embryonic stem cells in order to generate mice with conditional alleles. Basic Protocol 2 describes how to recover Cre transgenic mice from cryopreserved sperm using in vitro fertilization. Basic Protocol 3 describes a breeding strategy for obtaining tissue-specific knockouts. Finally, Basic Protocols 4 – 6 detail methods to validate the knockout mice using genomic DNA, reverse-transcription polymerase chain reaction (RT-PCR), quantitative reverse-transcription polymerase chain reaction (qRT-PCR), and Western blot analysis.

INTRODUCTION

The use of knockout mice to study gene function is a staple in toxicology research. Conditional knockout strains, in which gene function is altered in a cell/tissue-specific manner, are particularly powerful (Taylor et al., 2019). The Cre-loxP system is widely used for generating conditional knockouts (Papaioannou & Behringer, 2005). In this system, Cre recombinase, derived from the bacteriophage P1, catalyzes the recombination between two 34-bp loxP recognition sequences (Klos, 2004; Papaioannou & Behringer, 2005). To make mice with a conditional allele, loxP sites oriented in the same direction are introduced to the mouse genome to flank a critical exon or exons of a particular gene (these genes are called “floxed”). The expression of Cre recombinase in a specific cell type results in the deletion of the sequence between the two loxP sites. A schematic of the Cre-loxP system is shown in Figure 1.

Figure 1. Overview of the Cre-loxP System.

In the Cre-loxP system, inserted loxP sites flank the target region. Here, the loxP sites flank exon 2 within the targeted gene. When Cre, under the control of a specific promoter, is expressed, it binds to the loxP sites. Cre recombination occurs, knocking out the targeted region.

Conditional knockout mice are an ideal model to study organ-specific gene function and human disease. For example, to understand how loss of the manganese efflux transporter SLC30A10 induces neurological disease, we used the Cre-loxP system to generate full-body and tissue-specific Slc30a10 knockout mice. Full-body Slc30a10 knockout mice exhibited extremely elevated manganese in the brain, blood, and liver (Hutchens et al., 2017; Liu et al., 2017; Taylor et al., 2019). These knockouts were also hypothyroid, highlighting a previously unappreciated relationship between manganese toxicity and thyroid function (Hutchens et al., 2017; Liu et al., 2017; Taylor et al., 2019). Interestingly, brain-specific Slc30a10 knockouts had no change in brain manganese levels under basal conditions; while liver- and digestive system-specific knockouts showed moderate and extreme elevations in brain manganese, respectively (Taylor et al., 2019). Comparing the phenotypes of full-body and tissue-specific Slc30a10 knockout mice resulted in the unexpected discovery that brain manganese homeostasis is regulated by manganese efflux in the digestive system (Taylor et al., 2019). This application of the Cre-loxP system led to critical insight into the mechanisms of manganese homeostasis and manganese-induced neurological disease and underlines the power of using tissue-specific knockouts in research.

Here, we provide a detailed protocol for the generation and validation of tissue-specific knockout mice using the Cre-loxP system. Basic Protocol 1 describes the generation of mice containing loxP sites that flank a specific gene. Basic Protocol 2 describes how mice expressing the Cre transgene can be recovered. Basic Protocol 3 describes a breeding strategy for obtaining tissue-specific knockouts. Finally, Basic Protocols 4 – 6 detail four methods to validate the knockouts generated – 1) using genomic DNA, 2) using RT-PCR, 3) using qRT-PCR, and 4) using Western blot.

BASIC PROTOCOL 1: GENERATING MICE WITH CONDITIONAL ALLELES BY GENE TARGETING IN MOUSE EMBRYONIC STEM (ES) CELLS

Mice with a conditional allele, in which a critical exon(s) is flanked by loxP sites oriented in the same direction (floxed), can be made by gene targeting in mouse embryonic stem (ES) cells or by CRISPR-Cas9 genome editing. CRISPR-Cas9 can be faster and less labor intensive, but there are a number of limitations, including varying efficiencies and unintended mutations (Kosicki, Tomberg, & Bradley, 2018; Lanza et al., 2018). A discussion of the CRISPR-Cas9 system is provided in the Historical Background section of this manuscript. Here, we present only the gene targeting in ES cells approach. This approach involves four steps: 1) electroporation of a gene targeting vector into ES cells, 2) selection and picking of ES cell clones after electroporation, 3) identification of correctly targeted clones by Southern analysis, and 4) expansion of correctly targeted clones for microinjection. General considerations and strategies for designing gene targeting vectors to make mice with conditional alleles have been reviewed previously (Papaioannou & Behringer, 2005) and are not covered here. Targeting vectors can be created online and purchased commercially (Vector Builder, Cyagen). It is vital to design a Southern screening strategy to detect homologous recombination and identify single copy probes that lie outside of the homology arms of the targeting vector before electroporating the vector into cells. The design of the screening strategy should be coincident with the design of the targeting vector. Restriction enzymes sites at the locus and DNA fragments that may be suitable to use as single copy probes for Southern analysis can be identified using the University of Santa Cruz genome web browser. Although Southern analysis is generally more informative for identifying clones that are correctly targeted, an alternative protocol for identifying targeted clones by long-range PCR is also provided. The cell culture protocols described here can also be used to culture EUCOMM and KOMP C57BL/6N ES cell clones that have conditional alleles. EUCOMM and KOMP ES cell clones for a specific gene can be searched for at the International Mouse Strain Resource (IMSR). The IMSR links provide allele details, quality control information, and ordering information for the clones selected.

Materials

ES cells – V6.5 129/B6 hybrid mouse ES cells, (Novus Biologicals, NBP1–41162)

Feeder cells – SNL 76/7 STO cells (ATCC SCRC-1049)

Mitomycin C (Sigma, M0503)

Dulbecco’s Phospate Buffered Saline (DPBS) without Calcium, Magnesium (Hyclone SH30028.02)

Knockout DMEM (ThermoFisher, 10829018)

Fetal Bovine Serum, Defined (Hyclone, SH30070.03)

100X Pen-Strep (Gibco, 15140–122)

100X L-Glutamine (Gibco, 25030–081)

100X 2-Mercaptoethanol =10 mM 2-Mercaptoethanol (Sigma, M3148) in DPBS and filter sterilized

0.25% Trypsin, (Gibco, 25200–056)

0.1% Gelatin (Millipore, ES-006-B)

STO medium (see recipe in Reagents and Solutions)

ES cell medium (see recipe in Reagents and Solutions)

2x Freezing medium (see recipe in Reagents and Solutions)

Lysis Buffer (see recipe in Reagents and Solutions)

Hybridization Buffer (see recipe in Reagents and Solutions)

ES cell injection medium (see recipe in Reagents and Solutions)

G418 Sulfate (Geneticin, Gibco 11811–023)

Proteinase K (Roche, 03115879001)

LongAmp Taq 2x Master Mix (New England Biolabs, M0287S)

Plasticware: tissue culture-grade 10 cm dishes (Corning, 25020); 6 cm dishes (Falcon, 353004), 96-well flat bottom plates (Falcon, 353072); 4-well plates (Nunc, 179820); 12-well plates (Falcon, 353043); 5, 10, and 25 mL pipettes; autoclaved 200 μL pipette tips; reagent reservoirs; autoclaved Pasteur pipettes

2 mL cryogenic vials (Corning, 430488)

Blotting paper, 703 (VWR, 28298–020)

Gene Pulser Xcell™ Electroporation Systems with a capacitance extender (BioRad)

Gene Pulser cuvette, 0.4 cm electrode (BioRad, 165–2088)

8-channel aspirator (Corning, 4931)

20–200 μL 8-channel pipettor (VWR, 89079–948)

Owl Model A2 large gel system (Thermo Scientific)

UVT gel tray (Thermo Scientific, A2-UVT)

24 well 1.5 mm comb (Therm Scientific, A2–24D)

Hybridization bags (VWR, 95059–394)

Table top Impulse Sealer – 12 inch (Uline H-190)

Megaprime DNA labeling system (GE Healthcare, RPN1604)

Amersham Hybond-XL (GE Healthcare, RPN203S)

Carestream Biomax XAR film (Sigma, F5513)

Cell freezing container (Bel-Art, F18844–000)

Protocol 1.1 – ES cell electroporation

ES cells are expanded on a layer of SNL76/7 feeder cells, electroporated with the targeting vector, and selected with G418 Sulfate. The resulting colonies are picked, disaggregated with trypsin, and cultured using 96-well feeder plates. The 96-well plates are then split to make a plate for DNA isolation that is used for screening and a plate that is frozen for later clone retrieval.

Notes for working with ES cells.

To maintain their germline potential, mouse ES cells should be grown at high density on mitotically inactivated SNL76/7 feeder cells. The feeder cells provide Leukemia Inhibitory Factor (LIF) and other unknown factors that keep the cells in an undifferentiated state. ES cells need to be passaged approximately 1:6 every second or third day to prevent them from overgrowing and differentiating. During the trypsinization step, cell clumps must be disrupted by repeated pipetting to ensure a single cell suspension as cell clumps will differentiate. Cells should be fed every morning, and the medium should not be allowed to become yellow. Record the passage numbers of both ES cells and SNL76/7 cells. Do not use SNL76/7 cells after passage 20. The V6.5 (C57BL/6 × 129/Sv) ES cells are very robust, and the stocks that we use for electroporation are passage 26. If 129 strain ES cells are used instead of the hybrid V6.5 line, it is recommended to start with cells that have passage numbers in the low to mid-teens. It is also imperative to plan for the quantity of feeder plates needed for the experiment so there are enough SNL76/7 cells on hand to make the feeder plates. A confluent 10 cm plate contains approximately 7–8 × 10⁶ cells. Prepare feeder plates at least one day before using. Feeder plates are good for approximately two weeks, but they should be inspected before use to make sure the feeder layer is intact. Add new 1X glutamine to media after two weeks.

• 1
  Remove a vial of SNL76/7 cells (1 mL, 5 × 10⁶ cell/mL, passage number < 16) from liquid nitrogen storage and place the vial in a 37° C water bath for approximately 5 min.

  Please note that the caps of cryovials should be loosened immediately after removal from liquid nitrogen to let any liquid nitrogen that might have gotten into the vial to evaporate. Failure to loosen the cap may result in the tube exploding. A lab coat, cryo gloves, safety glasses, and a face shield should be worn when removing clones from liquid nitrogen storage.

• 2Transfer the thawed cells to a 15 mL tube containing 5 mL of STO medium to dilute out the cryoprotectant medium. Spin down the cells in a clinical centrifuge for 10 min at approximately 3,000 rpm.
• 3
  Remove the medium, re-suspend the cell pellet in 5 mL of STO medium, and transfer to a 10 cm gelatinized plate containing 7 mL of STO medium. When the cells become confluent in two to three days, passage the cells 1:5 to 5 × 10 cm gelatinized plates.

  Plates are gelatinized by pipetting enough 0.1 % gelatin solution to cover the plate. The gelatin is removed after 10–15 min.

• 4
  Make a 0.5 mg/mL mitomycin stock by adding 4 mL of DPBS to a bottle containing 2 mg mitomycin. Add mitomycin to the required amount of STO medium to a final concentration of 10 μg/mL.

  The mitomycin stock can be stored at 4° C for two weeks in its original dark vial. Mitomycin is a suspected carcinogen, so handle carefully and dispose of stocks and working solutions as chemical waste as directed by your institution’s chemical safety department.

• 5When the SNL76/7 plates are confluent, remove the medium from the SNL76/7 plates, and replace with 6 mL of STO medium containing mitomycin per 10 cm plate. Return the plates to the CO₂ incubator for two hours.
• 6Aspirate off the mitomycin-containing medium (collect it as chemical waste) and wash the plates two times with DPBS. Add 1 mL of trypsin to each plate and incubate at 37°C for 5–10 min. After incubation, add 5–10 mL of feeder medium (no mitomycin) to each plate to inactivate the trypsin and disaggregate the cells by pipetting up and down approximately ten times. For 5 × 10 cm plates, pool the cells between 2 × 50 mL tubes and centrifuge the tubes for 10 min at ~ 3,000 rpm. Remove the medium and wash the cell pellets twice with fresh STO medium to remove all traces of mitomycin. Re-suspend the cells in each tube in 10 mL of STO medium and count the number of cells in a 10 μL aliquot using a hemocytometer. Adjust the cell concentration to 3.5 × 10⁵ cells/mL. For electroporation, prepare 2 × 6 cm gelatin-coated feeder plates (4 mL of cells per plate) and 8 × 10 cm gelatin-coated feeder plate (12 mL of cells per plate). For this example, a total of 36 × 10⁶ mitomycin-treated cells would be required. A confluent 10 cm SNL76/7 plate contains 7–8 × 10⁶ cells, so approximately five SNL76/7 plates would be required to make the necessary feeder plates. Place the feeder plates in the incubator overnight to allow the mitomycin-treated cells to attach and recover.
• 7Thaw and re-suspend a vial of V6.5 ES cells as described above for SNL76/7 cells. Re-suspend the cells in 4 mL ES medium and plate to a 6 cm feeder plate. The following morning, feed the cells. It will take two to three days for the cells to become approximately 70% confluent. The ES cells can then be passaged 1:2 to a 10 cm feeder plate.
• 8The next morning, feed the ES cells. In the afternoon (~ 4 hours after feeding), trypsinize the ES cells for 10 minutes and gently disaggregate the cells to make a single cell suspension. Spin down the cells and re-suspend in 10 mL DPBS. Count a 10 μL aliquot of cells using a hemocytometer. Adjust the cell concentration to 11 × 10⁶ cell/mL in DPBS. Place 0.9 mL of cells into a Gene Pulser cuvette and add 25 μL of the linearized targeting vector (1 mg/mL). Place the cuvette in the electroporation pod and electroporate at 230 V and 500 μF. Let the cells recover on ice for 5 min. Replace the medium on 6 × 10 cm feeder plates with ES medium and aliquot the electroporated cells between the six feeder plates.

Protocol 1.2 – Selection and picking ES cell clones

• 9Start selection with G418 approximately 24 hours after electroporation. For V6.5 ES cells, use 200 μg/mL G418 in ES medium. For other ES cell lines, test various concentrations of G418 to determine the appropriate concentration for selection. Feed the cells daily for the first four to five days. After this initial period, cells can be fed every day or every other day, depending on the number of clones on the plate. Clones can be picked 8–10 days after electroporation. Maintain the G418 selection until picking the clones.
• 10To pick the ES clones, rinse the plates twice with DPBS and cover with 10 mL of DPBS. Colonies are picked using a Gilson PIPETMAN set to 2 μl and placed in individual wells of a 96-well plate containing 50 μl of trypsin per well. Pick clones under a stereomicroscope set at low power to identify clones that have a rounded morphology. Avoid clones with a flat phenotype; the rounded morphology should be visible by eye. Do this outside of the cell culture hood in a quiet area of the lab to prevent contamination. Once a full 96-well plate has been picked, which takes approximately 30 min, the 96-well plate of ES cell clones is placed in the incubator for 10 min. Use a multichannel pipettor, to add 50 μl of ES medium to each well, and break clones up by pipetting up and down approximately 20 times using the multichannel pipettor. Transfer the cells from each well to another 96-well feeder plate. To make 96-well feeder plates, use 150 μL of 3.5 × 10⁵ mitomycin-treated cells per well. The steps in this process are also provided in Figure 2.
• 11Feed the ES cells daily for three to four days until the wells become almost confluent.
• 12When nearly confluent, aspirate off the medium, wash twice with DPBS, add 50 μL of trypsin per well and incubate for 10 min at 37° C. Then add 50 μL of ES cell medium and disaggregate the cells. Transfer 50 μL of cells to a gelatin-coated 96-well plate and culture for four to five days. This plate will be used for DNA isolation.
• 13To the remaining 50 μL of cells, add 50 μL of 2x Freezing medium and mix several times. Wrap the freeze plate in parafilm and place in a food storage bag. Put the plates in a small Styrofoam box and place the box in a −80° C freezer.

Figure 2. Steps in picking ES cell clones.

Please see text (Basic Protocol 1.2 Step 10) for details of the four steps pictures.

Protocol 1.3 – DNA isolation and Southern analysis

DNA from the ES cell clones is isolated by a simple extraction technique and digested with a restriction enzyme directly in the 96-well plate (Ramirez-Solis et al. 1992). EcoRI, EcoRV, BamHI, BglII, PstI, Asp718, and SstI give complete digestions using this isolation procedure. Following restriction digestion, the DNA is run on a gel containing four combs of 24 wells each, so that one 96-well plate equals one gel. The DNA is then transferred to a nylon membrane by Southern blotting and hybridized with radioactively labeled 5’ and 3’ single copy probes that lie outside the arms of homology of the vector. Blots are then washed and exposed to X-ray film to detect clones that have undergone targeting.

• 14Wash the 96-well plates twice with 100 μL DPBS per well and then add 50 μL per well of freshly prepared lysis buffer. Seal the plates with parafilm and place them in a Tupperware-type container containing a wet paper towel to provide humidity. Place the container in a 60° C water bath overnight.
• 15Precipitate the ES cell DNA by adding 100 μL of a NaCl/ethanol mix (150 μL 5 M NaCl and 10 mL100% ethanol) to each well. Let stand for 30 minutes. The precipitated DNA will form a white meshwork in the well. Slowly pour off the ethanol, keeping the DNA in the well. Gently rinse the plates 5–6 times with 70% ethanol and air dry.
• 16Digest the DNA by adding 30 μL of restriction enzyme mixture containing 30 units of enzyme. Seal the plates with parafilm and incubate overnight in a sealed humidified container.
• 17Run the DNA samples on an appropriate percentage agarose gel containing 24 wells across and four rows of wells (20 × 24 cm gel tray). Run the gel for several hours at 60 – 70 volts until the bromophenol blue of the loading dye migrates to just before the next well set. Photograph the gel.
• 18
  Prepare an apparatus for Southern blotting by completing the following (shown in Figure 3):

  1. Place three large sponges in a large glass baking dish.
  2. Cut four pieces of blotting paper the size of the gel.
  3. Cut one piece of the Hybond-XL blotting membrane the size of the gel and pre-wet the membrane by placing it in a tray of distilled water.
  4. Flood the sponges in the baking dish with 1 L of 0.4 N NaOH.
  5. Place two sheets of blotting paper on top of the sponges.
  6. Flood with more 0.4 N NaOH and smooth out any bubbles in the blotting paper by rolling a glass pipette across the surface.
  7. In another glass baking dish, nick the DNA in the gel by incubating the gel in 0.25 M HCl (~ 500 mL) until the bromophenol blue dye band starts to turn yellow (typically about 7 minutes, but this depends on the thickness of the gel).

     The nicking step aids in the transfer of high molecular weight DNA out of the agarose gel.

  8. Carefully pour off the HCl solution and rinse the gel several times with tap water. Place the gel on the blotting paper-covered sponges.
  9. Flood the gel with more 0.4 N NaOH and place the Hybond-XL blotting membrane over the gel. Nick the bottom right corner of the gel and the membrane for orientation purposes.
  10. Flood again with 0.4 N NaOH and place the remaining two blotting papers on top of the Hybond-XL membrane.
  11. Add a 4-inch stack of paper towels on top of the blotting paper. Compress the paper towels lightly by adding a glass plate or the acrylic gel tray. Do not add heavy items as this may adversely affect the transfer. Let the transfer proceed for 4 hours. The 0.4 N NaOH transfer solution denatures the DNA so that it can bind the Hybond-XL membrane.
  12. Disassemble the transfer setup and rinse the Hybond-XL membrane twice with 2x SSC in a baking dish or tray. The bromophenol dye band transferred to the blotting membrane will turn blue again when the membrane is neutralized. Let the membrane air dry. For easy handling during hybridization and washing, the membrane can be labeled with a pencil and cut into four strips representing each well tier, and then placed in a hybridization bag until they are used.
• 19Label 100–200 ng of the 5’ and 3’ single copy hybridization probes using the MegaPrime labeling kit and 32P-dCTP (NEN BLU013H). Remove unincorporated 32P-dCTP by spinning the reaction through a G50 Sephadex syringe column. Count a 1 μL aliquot of the probe in a scintillation counter. Probes should have a specific activity of 0.5 −1 × 10⁹ counts per minute.
• 20Pre-hybridize the Hybond-XL strips in 50 mL of hybridization buffer in a heat-sealed hybridization bag. Pre-hybridize the membranes for 1 hour at 42° C in a shaking water bath. Pour out the hybridization buffer and add 20 mL of fresh hybridization buffer. Denature the probe by heating it to 95° C in a PCR machine for 5 min. Place the denatured probe on ice and add it to the hybridization buffer. Smooth out any air bubbles by carefully rolling a glass pipette across the bag. Seal the bag and place it in a shaking water bath at 42° C and hybridize for 24 hours. The hybridization bag can be weighted down using a glass plate or small weights placed on the edges of the bag.
• 21
  Wash the blots

  1. Pour off the radioactive hybridization solution and dispose as directed by your institution’s safety department.
  2. Transfer the blots into a Tupperware-type container with 500 mL of 2x SSC/0.1 % SDS and rinse once.
  3. Transfer the blots to a second container of the same solution and wash for 30 min at 65° C with gentle shaking. Dispose of the radioactive hybridization bag and your gloves as radioactive solid waste for disposal by your safety department.

     Monitor all work areas and your person with a GM counter for potential contamination with radioactivity.

  4. Wash the blots twice in 0.1X SSC/0.1 % SDS for 30 min each time at 65° C.
  5. Rinse the blots with 0.1X SSC and air dry the blots until just damp.
  6. Wrap the blot strips neatly in plastic wrap and orient the strips according to their position on the original gel. Place in an X-ray film cassette with an intensifying screen and expose to Biomax XAR X-ray film for at least 24 hours.

Figure 3. Steps in setting up Southern blot for ES cell clone screening.

A) Large sponges are placed in a glass baking dish and flooded with 0.4N NaOH. B) Two strips of blotting paper cut to the size of the gel are placed on the sponges and flooded with more 0.4N NaOH. A 10 ml pipette is used to roll out any air bubbles beneath the blotting paper. C) Following treatment with 0.25 M HCl, the agarose gel is placed on the blotting paper and flooded with more 0.4N NaOH and then covered with the nylon blotting membrane. More 0.4N NaOH is poured over the membrane. The pipette is used to roll out any air bubbles between the gel and the membrane that would impede DNA transfer to the membrane. Note the nick in the gel and the membrane in the bottom right hand corner for orientation (arrowhead). D) Two more strips of blotting paper are placed on top of the nylon membrane. Note the addition of strips of parafilm (arrow) around the sides of the set up to prevent short-circuiting of 0.4N NaOH transfer buffer between the sponges and the paper towels to be added. E) A stack of hand paper towels is added. Here for illustrative purposes only one stack of towels is shown. F) A light weight, here an agarose gel casting tray, is placed on top of the paper towels.

Protocol 1.4. – Clone expansion and preparation of ES cells for microinjection

Targeted clones are thawed and gradually expanded on consecutively larger feeder plates. Clones are fed daily with ES cell media and passaged when approximately 70% confluent. Cells from a 6 cm plate are frozen in cryovials and are stored in liquid nitrogen before microinjection. For V6.5 ES cell clones, expand three to six targeted clones and make at least three vials of frozen cells per clone. Begin by injecting three targeted clones per project.

• 22Remove a 96-well plate containing targeted clones from the Styrofoam box at −80° C and place in the CO₂ incubator for 15–20 min or until the clones are thawed.
• 23Transfer the cell contents, ~100 μL, from the appropriate well to a 4-well feeder plate containing 1 mL of ES medium.
• 24Feed the cells the next morning to remove the DMSO and oil.
• 25
  Feed the cells daily and passage all the cells to a 12-well feeder plate, and when 70% confluent passage 1:2 to two 6 cm feeder dishes.

  Do not let the cells overgrow, and do not passage the cells too sparsely.

• 26
  Trypsinize the cells from the 6 cm plate and count the cells with a hemocytometer. Adjust the cell concentration to 10 × 10⁶ cells/mL and add an equal volume of 2X Freezing medium.

  Make at least three vials of cells per clone. Place the vials in a cell freezing container and store at −80° C for 24 hours. Transfer vials to liquid nitrogen for long-term storage.

• 27For microinjection, remove a vial of cells from liquid nitrogen and thaw to a 6 cm feeder plate. Feed the cells daily and passage two to three days later to a new 6 cm feeder plate so that the dish will be approximately 60 – 70% confluent after two days. Feed the cells 4 hours before injection.
• 28
  To prep cells for injection, trypsinize the cells for 7–10 min, and disaggregate the cells into a single cell suspension.

  If a second injection is needed, in two days, passage the cells approximately 1:6 to a new 6 cm feeder plate. Take the remaining cells and place them in a 15 mL tube and centrifuge for 5–10 min at approximately 3,000 rpm. Gently re-suspend the cell pellet in 2–3 mL of ES cell injection medium and place the tube on ice for injection.

• 29Place approximately 200 – 300 μL of cells into an injection chamber containing ES cell injection medium. For V6.5 cells, inject 10–12 cells into each C57BL/6N blastocyst and 12–15 ES cells for 129 and C57BL/6 ES cell clones. Inject approximately 30 blastocysts for each V6.5 or 129 clone and 40–50 blastocysts for KOMP and EUCOMM C57BL/6N ES cell clones. Use C57BL/6 albino strain blastocysts when injecting C57BL/6N KOMP and EUCOMM clones.
• 30Transfer six to seven injected blastocysts to the oviduct or uterine horn per side for each recipient mouse. Potential chimeric mice are born 17–19 days later.
• 31
  Pick three to four high percentage chimeric male mice based on coat color for each clone and breed the males with C57BL/6N females to test for germline transmission.

  For V6.5 or 129 ES cell clones, if germline transmission has occurred, one or more pups will have an agouti coat. Genotype the agouti pups to identify mice that are heterozygous for the mutation. Chimeric males generated using EUCOMM or KOMP C57BL/6 JM8A3 cells in which the agouti mutation has been corrected can also be crossed with C57BL/6N female mice to test for germline transmission by observing the coat color. For C57BL/6N JM8 clones, chimeras can be mated with C57BL/6 albino strain mice to identify germline transmitters. As all the commercial C57BL/6 albino strains are J substrain, and the EUCOMM and KOMP ES cells are N substrain, breed the identified germline transmitters with C57BL/6N females, and genotype all the pups to identify heterozygous pups.

Alternate Protocol 1: SCREENING ES CELL CLONES BY LONG-RANGE PCR:

As an alternative to Southern analysis (Basic Protocol 1.3), ES cell clones can be screened by long-range PCR to detect clones that have undergone homologous recombination. DNA for PCR is prepared using the same protocol as for Southern analysis (Basic Protocol 1.3, Steps 14 – 15). Clones with a targeted allele are identified using a primer to a unique cassette sequence in the targeting vector, such as neo, and a second primer located outside the region of homology arm of the targeting vector. Sequence information for designing PCR strategies and primers can be obtained using the UCSC or Ensembl genome browsers.

This protocol is based on the LongAmp polymerase from New England Biolabs. Follow the manufacturer’s recommendations if using a different polymerase for PCR.

Additional Materials

LongAmp Taq 2x Master Mix (New England Biolabs, M0287S)

1. Re-suspend the DNA in the 96-well plate with 50–100 μL of DNAse/RNAse-free water or TE buffer per well.

2. Add 1 μL of ES cell DNA to 9 μL of Long Amp PCR cocktail for each 5’ and 3’ PCR reaction.

3. Run the PCR. For amplifying DNA bands up to 4 kb use the following:

   1. 95° C for 2 min.
   2. 35 cycles of 95° C for 30 sec., 60° C for 15 sec, and 65° C for 3 min.
   3. 5 min. extension at 65° C.

4. Run 5 μL of the PCR product using the same gel set up as for Southern analysis (4 × 24 well combs).

BASIC PROTOCOL 2: RECOVERY OF CRE TRANSGENIC LINES BY IN VITRO FERTILIZATION (IVF) USING CRYOPRESERVED SPERM

The second critical step in generating a tissue-specific knockout mouse using the Cre-loxP system is to obtain the desired Cre expressing mouse. Tissue specificity is established by the promoter controlling the expression of Cre recombinase. For example, the Sox2-Cre strain can be used to generate global knockouts, while the Nestin- and Albumin-Cre strains can be used to generate brain- and liver-specific knockouts, respectively (Hutchens et al., 2017; Liu et al., 2017; Taylor et al., 2019). In the Nestin-Cre strain, Cre expression is under the control of the Nestin promoter, which is expressed in neuronal and glial precursors (Tronche et al., 1999). Conversely, in the Albumin-Cre strain, Cre activity is under the control of the Albumin promoter, which is expressed in hepatocytes (Postic et al., 1998). Another important factor in choosing a Cre strain is the developmental day on which Cre expression begins. For example, in the Nestin-Cre strain, recombinase activity is observed by embryonic day 11 (Tronche et al., 1999). Thus, beginning embryonic day 11, cells derived from neural cells expressing Cre will have the gene deleted. There are also tamoxifen inducible Cre strains, in which Cre activity is induced following exposure to tamoxifen (El Marjou et al., 2004; Feil et al., 1996). This provides temporal control over full-body or organ-specific Cre recombination. Lastly, another factor to consider when selecting a Cre strain is any phenotype associated with that strain that may be related to the transgene integration site (Goodwin et al., 2019). It is essential to be aware of how strain-specific phenotypes may influence experimental results.

Many Cre strains are readily available through mouse repositories. These repositories often list original papers describing the strain, the expression pattern of the recombination, any strain-associated phenotypes, and recommendations for control animals. If the live animal is available, the desired Cre strain can be ordered and bred. If the live animal is not available, the transgenic line must be recovered (detailed below).

Sperm cryopreservation has become the standard method to archive transgenic mouse lines. Straws or vials containing aliquots of cryopreserved sperm can be shipped from repositories or core facilities to university core labs and the animals recovered by in vitro fertilization (IVF). This protocol is based on the IVF protocol developed at the Center for Animal Resources and Development (CARD) at Kumamoto University (Takeo & Nakagata, 2011). A kit based on the CARD IVF protocol is also available from Cosmo Bio Co., Ltd. It is critical to adhere to the given times when performing the IVF steps. Additionally, the quality of the paraffin oil is vital for successful IVF. The oil should be tested before using in IVF by culturing 1-cell embryos overnight under the oil to determine the percentage of embryos that develop to the 2-cell stage. If the oil is good, nearly all the embryos will advance to the 2-cell stage.

Materials

FERTIUP PM-CARD MEDIUM set (Cosmo Bio Co., Ltd. KYD-006-EX)

Pregnant Mare Serum Gonadotrophin (PMSG) hormone (ProSpec, HOR-272)

Human Chorionic Gonadotrophin (HCG) hormone (Sigma, 1063)

Water – embryo tested (Sigma, W1503)

HTF medium high calcium with BSA, (Cytospring, mH0114)

Paraffin oil (Nacalai USA, 2613785)

Pipette tips

35 cm plastic dishes

Water bath set at 37° C

Humidified incubator

Micropipettes for handling embryos

1. Prepare PMSG and HCG aliquots for superovulation by reconstituting the hormone with water to a concentration of 1000 IU/mL. Aliquot 50 μL of hormone per tube and store the aliquots at −80° C. Immediately before superovulation, thaw the hormone and add 950 μL of sterile saline to make a 50 IU/mL injection stock.

2. Superovulate female mice by injecting 5 IU (100 μL) PMSG at 6:00 pm on Day −3 and 5 IU HCG at 6 pm on Day −1.

   For each experiment use six female mice, approximately 8–12 weeks of age.

3. On Day −1, prepare a dish with 4 × 150 μL drops of HTF medium covered with paraffin oil and place in a CO₂ incubator. Also, place a 50 mL tube of water in a beaker in the incubator to be used for thawing sperm the following day.

4. At 7:30 am on Day 0, prepare a plate with 90 μL drop of FERTIUP, cover with paraffin oil, and place in the incubator. Make a second dish with 90 μL drop of CARD medium; cover with paraffin oil and place in the incubator. Follow the kit instructions for preparing the individual FERTIUP and CARD reagents.

5. At 8:00 am, euthanize a female mouse by cervical dislocation. Work with one mouse at a time. Remove the oviducts from the mouse and place into the paraffin oil alongside the CARD medium. With forceps tear the oviducts to release the oocyte-containing cumulus masses into the oil. Drag the cumulus masses into the CARD medium drop. Figure 4 provides an image of this process.

   It is crucial to work as quickly as possible. The entire process should take less than a minute.

6. Repeat the oviduct collection steps with the remaining mice. Return the dish with the oocytes to the incubator.

7. At 8:30 am, remove the straw containing the sperm sample from liquid nitrogen. Immerse the straw in a 50 mL beaker partially filled with water pre-warmed to 37° C.

8. Return the tube to the 37° C incubator and leave undisturbed for 10 min.

9. After 10 min, remove the straw and dry the outside of the straw with a paper towel. Cut the ends of the straw and carefully expel only the sperm aliquot (~ 10 μL) into 90 μL drop of FERTIUP. Incubate the sperm for 30 min at 37° C in the CO₂ incubator.

10. At 9:00 am, remove 10 μL of the thawed sperm from FERTIUP medium using a 200 μL pipette tip cut at an angle to give a larger bore. Remove the sperm from the edge of the drop, to collect the more highly motile sperm. Add the 10 μL of sperm solution to oocytes in CARD medium and incubate for 3 hours in the CO₂ incubator.

11. After the incubation period, wash the oocytes through three drops of the HTF medium. Move the oocytes to the fourth HTF drop and culture them overnight.

12. The next morning, count the number of two-cell embryos and transfer them to the oviducts of pseudopregnant recipient mice.

Figure 4. Isolation of mouse oocytes for IVF.

A) The ovary with the attached oviduct is shown towards the bottom of the image. The oocytes are removed from the ampulla of the oviduct by tearing the oviduct wall with forceps. The released oocytes surrounded by cumulus mass cells are then dragged into the HTF drop. B) Higher magnification image of isolated oocytes surrounded by cumulus mass cells in the HTF droplet.

Note: The protocol described above is for sperm cryopreserved using the CARD method, which is widely used by mouse repositories and University transgenic core labs because of its simplicity and efficiency. For working with sperm prepared by older methods, use a slight modification of the protocol described by CARD (Nakagata et al., 2014). Transfer the thawed sperm to a 1.5 mL microtube using a wide bore pipette tip and centrifuge the sample at 300 rpm for 5 min. Remove the supernatant and re-suspend the sperm sample to approximately 100 μL using FERTIUP medium. Place the sperm solution in 35 mm dish and top with oil that has been pre-equilibrated in the CO₂ incubator for at least 12 hours. Incubate the sperm suspension for 30 min in the CO₂ incubator and add it to the oocytes as described in step 8 above.

BASIC PROTOCOL 3: BREEDING STRATEGY TO GENERATE TISSUE-SPECIFIC KNOCKOUT MICE

Once both the floxed mouse and Cre expressing mouse are obtained, two rounds of breeding must occur in order to produce knockout mice. Generating a tissue-specific knockout mouse requires accurate genotyping of animals maintained for experimental crosses. Support Protocol 1 describes a general genotyping strategy for tissue-specific knockout mice.

Materials

Homozygous floxed mouse (loxP sites flanking target region)

Cre strain

Ear punching tool for mice

1. Breed the Cre strain with a homozygous floxed mouse. Genotype progeny.

   This breeding pair will produce animals that are heterozygous for the loxP sites, with or without Cre. Note that this first breeding pair will not produce knockout mice.

2. Breed a homozygous floxed mouse with a mouse heterozygous for the loxP sites and expressing Cre. Genotype progeny.

   This breeding pair will produce mice that are heterozygous or homozygous for the loxP sites, with or without Cre. The mice homozygous for the loxP sites and expressing Cre are the knockout mice.

Support Protocol 1: Genotyping of stock mice

The protocol for genotyping is very similar to the protocol used for validating a knockout using genomic DNA (Basic Protocol 4). Please see Basic Protocol 4 for the step by step protocol for DNA extraction, PCR, and gel electrophoresis. The key differences are the tissue used for DNA extraction and the primers. Tissue collected from ear punches (or tail snips) should be used for the DNA extraction. For the DNA extraction only 50 μL of DNA extraction reagent and DNA stabilization buffer is needed as ear punches are much smaller than tissue samples. Because ear punches or tail snips may not be the target tissue for Cre recombination, there is no need to conduct the PCR for the knockout allele as it should not be present in non-target tissue. Instead a PCR to detect the loxP sites and a PCR to detect Cre must be performed. Considerations when designing primers are also detailed under Support Protocol 2. Figure 5 provides a schematic for Primer Set 1 and Primer Set 2 placements.

Figure 5. Primer placement for genotyping.

The forward and reverse primers for Primer Set 1 (P1-FWD and P1-REV, respectively should be placed around the 5’ loxP site. The forward and reverse primers for Primer Set 2 (P2-FWD and P2-REV, respectively should be placed around the 3’ loxP site. Since each loxP site is 34 bps, the resulting PCR product from a floxed allele should be 34 bps longer than the PCR product from a wildtype allele.

Primer Set 1: Detection of the 5’ loxP site.

The forward primer should begin just before the start of the 5’ loxP site. The reverse primer should begin just after the 5’ loxP site. Because each loxP site is 34 bp, the total PCR product when loxP sites are present should be longer than the wildtype allele, which has no loxP sites (Klos, 2004). PCR product from heterozygous mice will show 2 bands, while product from homozygous mice will show 1 longer band.

Primer Set 2: Detection of the 3’ loxP site.

The forward primer should begin just before the start of the 3’ loxP site. The reverse primer should begin just after the 3’ loxP site. The total PCR product when loxP sites are present should be longer than the wildtype allele, which has no loxP sites. PCR product from heterozygous mice will show 2 bands, while product from homozygous mice will show 1 longer band.

Primer Set 3: Detection of Cre.

Forward and reverse primers can be designed to probe any part of the Cre transgene. This primer set will produce a single band when Cre expression is present and no band if there is no Cre.

BASIC PROTOCOL 4: IDENTIFICATION OF EXON DELETION IN TARGET TISSUE

When the Cre-loxP system is used to generate tissue-specific knockout mice, Cre expression should only be found in the targeted tissue(s). For example, if Nestin-Cre is used to generate a pan-neuronal/glial knockout, the Cre should be expressed in neurons and specified glial cells (oligodendrocytes and astrocytes), but not in other tissue, such as the liver. Additionally, the recombined allele should only be expressed the target tissue of mice homozygous for the floxed allele and expressing Cre. These limitations on where the Cre should be expressed and active allow for the use of genomic DNA from target and off-target tissues to validate a successful and specific knockout of the desired gene.

Broadly, the procedure for validating a knockout using genomic DNA involves designing appropriate primers (Support Protocol 2), extracting DNA from necessary tissue, conducting polymerase chain reaction (PCR) using designed primers, and electrophoresis of the PCR products on an agarose gel. The following protocol was used to validate knockout out the manganese efflux transporter Slc30a10 in specific tissue in mice (Taylor et al., 2019).

Tissue collection for this protocol should be done with RNAse- and DNAse-free materials. Once the tissue is extracted, it may be placed in a 1.5 mL microcentrifuge tube. If tissue will not be used immediately, flash freeze tissue in liquid nitrogen and store at −80°C until ready for use. The protocol for DNA extraction and PCR are based on information provided by Quantabio. Please see reagent/kit manuals for instructions if not using the same as those listed below.

Materials

Fresh/Fresh frozen tissue (one sample of the target tissue and one sample of off-tissue for both a knockout and control mouse, ~50 mg each)

DNAse/RNAse removal solution

DNA extraction reagent (Quantabio, 84158)

DNA stabilization buffer (Quantabio, 84159)

Ice bucket with ice

1.5 mL microcentrifuge tube

DNAse/RNAse-free water

2X PCR SuperMix (Quantabio, 95136)

Primers (100 μM, see Support Protocol 2)

PCR tubes (.2 mL)

Agarose

1X Tris-Acetate Ethylenediaminetetraacetic acid (EDTA) buffer (1X TAE) (see recipe in Reagents and Solutions)

DNA ladder of appropriate size (Thermo Scientific, SM0244 (100 bp), SM0314 (1 kb))

Heat block (able to reach 95°C)

Microcentrifuge (mini-centrifuge able to fit 1.5 mL tube)

Vortex mixer

Mini-centrifuge (able to fit PCR tubes)

PCR machine

Microwave

Electrophoresis power supply (PowerPac Universal Power Supply, Bio-Rad)

Submerged horizontal electrophoresis cell (Wide Mini-Sub Cell GT Cell, Bio-Rad)

Agarose gel casting kit (Wide Mini Handcasting Kit, Bio-Rad)

Gel Imaging System (GE Amersham Imager 600QC)

Protocol 4.1 – DNA Extraction

• 1Set heat block to 95°C.
• 2Add 100 μL of DNA extraction reagent to each tissue sample. Ensure each sample is clearly labeled.
• 3Briefly centrifuge samples at 12,000 rpm for 30 s to ensure the tissue is submerged in the extraction reagent.
• 4Place the samples on the heat block at 95°C for 30 min.
• 5After 30 min on the heat block, centrifuge samples at 12,000 rpm for 30 s.
• 6Add 100 μL of DNA stabilization buffer to each tissue sample and place on ice (or store at −20°C until ready to proceed to Step 7).

Protocol 4.2 – PCR

• 7Place PCR SuperMix and primers on ice and allow to thaw.
• 8Make a master PCR mix for each primer set (see recipe in Reagents and Solutions).
• 9Aliquot 20 μL of each master PCR mix for each sample in a PCR tube (i.e. if there are three different master PCR mixes, aliquot one of each per sample, so that each sample has 3 PCR tubes with three different master PCR mixes in them). Keep PCR tubes on ice.
• 10For each DNA sample and PCR master mix, add 2 μL of DNA sample to the 20 μL of master PCR mix. Pipette to mix.
• 11Briefly, centrifuge PCR tubes in the mini-centrifuge.
• 12
  Run necessary PCR conditions in the PCR machine. (PCR conditions are based on the primers and length of the PCR product. The annealing temperature should be set to ~5°C less than the lowest T_m of the primers. For the AccuStart II PCR SuperMix, 1 min per kb of PCR product is recommended for the extension time). The following was used to detect PCR products of ~250–500 bp:

  1. 94°C for 3 min
  2. 30 cycles of 94°C for 30 s, 60° for 30s, and 72°C for 1 min
  3. 72°C for 5 min
  4. 4°C for ∞

Protocol 4.3 Gel Electrophoresis

• 13
  When the PCR has approximately 30 min remaining, make the agarose gel for gel electrophoresis: Measure 2 g of agarose in a microwaveable flask. Add 100 mL of 1X TEA (see recipe in Reagents and Solutions).

  This specific protocol calls for a 2% agarose gel, however, the percent of agarose is based on the PCR product size.

• 14
  Microwave the 1X TAE + agarose for ~45 s. Swirl to mix. Microwave for ~15 s. Gently swirl to mix. Microwave for ~30 s and swirl to mix. When the agarose is fully dissolved in the TAE, the solution will be clear. Microwave an additional 20 s if the solution is not clear. Repeat as necessary until the agarose is fully dissolved.

  As the solution heats, it will boil. Do not let the mixture boil over.

• 15
  Remove the TAE + agarose mixture from the microwave and allow it to cool until it can be touched by hand. As the TAE + agarose solution is cooling, put together the agarose gel cast kit. Be sure to place the combs in the casting tray.

  The comb size is based on the total number of samples to be run. Ensure there are enough wells for each sample and the DNA ladder(s).

• 16Add ethidium bromide to the TAE + agarose mixture so that that final concentration is ~.5 – .75 μL/mL (i.e. add ~15 μL of a 5 mg/mL stock of ethidium bromide to 100 mL solution). Swirl to mix.
• 17
  Pour the mixture into the gel cast. Use a pipette tip to remove/move any bubbles. There should be no bubbles in near the wells or near where sample will be running. Allow at least 20 min for the gel to fully harden.

  If the PCR has not finished running, leave the gel at this step until the PCR is done.

• 18
  Once the gel has hardened, remove the gel casting tray with the gel and combs from the casting gate, and place in the horizontal electrophoresis cell.

  In the electrophoresis cell, black is negatively charged and red is positively charged. DNA is negatively charged, so the wells need to be placed closer to the negatively charged part of the electrophoresis cell (black). This allows the DNA to flow towards the positive (red).

• 19Fill the electrophoresis cell with 1X TAE, and carefully remove the combs from the gel.
• 20Add ~5 μL of DNA ladder to one well in each set of wells (i.e. if there are two sets of wells in the gel, there should be a well containing DNA ladder for each set). Add ~10 μL of sample to the well (one sample per well).
• 21Place the top on the electrophoresis cell. Be sure to match black to black and red to red. Connect the top to the electrophoresis power supply, again connecting black to black and red to red.
• 22Run the gel at a constant 100 V. If the gel is running correctly, the dye and yellow front should be moving away from the wells. Run the gel until the yellow front is ~3/4 of the way down the gel (if there are two sets of the wells in one gel, run the gel until the yellow front from the top row of wells is ~3/4 of the way to the bottom wells).
• 23Once the run is complete, remove the gel tray, and place the gel tray and gel (or just the gel) in the gel imager (Place the gel on the imaging tray appropriate for fluorescent UV imaging). Image the gel using the UV light setting. Save image.
• 24Dispose of the gel in a waste container designated for items containing ethidium bromide.

Support Protocol 2: Designing primers for validation using genomic DNA

Validation using genomic DNA requires assessing the presence of the knockout allele and the presence of Cre. Thus, two separate primer sets are needed. The first primer set aims to detect if recombination occurred, indicating knockout of the targeted gene. The second primer set aims to detect if the Cre is expressed. A primer set consists of the forward primer and the reverse primer. The forward primer consists of the exact DNA base pair sequence at the beginning of the desired sequence region. The reverse primer consists of the reverse complement of the base pair sequence at the end of the desired region. Primers should be ~15–20 base pairs in length, and the sequence should be unique. Below are recommendations for designing each primer set needed to validate knockouts using genomic DNA. Once the primer sequences are determined, primers can be ordered through Sigma-Aldrich or other online service.

Primer Set 1: Detection of recombination.

The forward primer should end at least 50 – 100 bases before the start of the 5’ loxP site. The reverse primer should begin 50 – 100 bases after the 3’ loxP site. With recombination, the 3’ loxP site and sequence between the loxP sites are removed. Figure 6 shows primer placements with and without recombination. The primers should be designed such that with recombination, the PCR product is ~250–500 bps, but without recombination the product size is greater than 1 kb (which is too large for the PCR conditions set). Thus, with this primer set, a PCR product from a knockout allele will produce a single band, while PCR product from a wildtype allele will produce no band.

Figure 6. Primer placement for detecting the recombination in genomic DNA.

The forward primer should be placed just before the 5’ loxP site, and the reverse primer should be placed just after the 3’ loxP site. Without recombination, the PCR product size is too large for the PCR conditions set. However, with recombination, the region between the loxP sites is excised, making the PCR product short enough for amplification using the PCR conditions described.

Primer Set 2: Detection of Cre.

Forward and reverse primers can be designed to probe any part of the Cre transgene. This primer set will produce a single band when Cre expression is present and no band if there is no Cre.

BASIC PROTOCOL 5: VALIDATION OF TRANSCRIPT REMOVAL USING RT-PCR AND qRT-PCR

In tissue-specific knockouts generated using the Cre-loxP system, Cre expression should be specific to the target tissue, and Cre activity should be specific to the floxed allele. Thus, in a tissue-specific knockout model, only target tissue should lack expression of the floxed allele. The ideal method to validate a tissue-specific knockout is to perform immunoblots to ensure that the targeted protein is depleted not in organs that express Cre. However, if antibodies are not available to detect the target protein, RT-PCR and qRT-PCR may be used to assess whether the knockout was both successful and specific.

RT-PCR involves extracting RNA from a tissue sample and reverse transcribing that RNA into cDNA. That cDNA product is amplified using PCR, and the PCR product is separated and visualized using gel electrophoresis, as described in Basic Protocol 4. If a knockout is both successful and specific, the targeted gene should be absent in target tissue, but present in all other tissue. Additionally, expression of a control gene ubiquitously expressed should also be analyzed to compare expression levels. The following protocol was used to assess expression of SLC30A10 and the mouse housekeeping gene, 18S, in tissue-specific Slc30a10 knockout mice (Taylor et al., 2019).

Tissue collection should be completed with RNAse and DNAse-free materials. Once extracted, tissue should be placed in a 1.5 mL microcentrifuge tube and immediately flash frozen in liquid nitrogen in order to preserve the quality of RNA. Tissue can then be processed for RNA extraction (detailed below) or stored at −80°C until ready for use. This protocol is based on the PureLink™ RNA Mini Kit and the cDNA synthesis reagents from Thermo Fischer Scientific. Please see kit(s) instruction manuals if not using the same materials described here.

Materials

Fresh frozen tissue (one sample of the target tissue and one sample of off-tissue for at least three knockout and three control mice, ~50 mg each)

DNAse/RNAse removal solution

Delicate task wipes

2-Mercaptoethanol (BME)

Pellet pestle for each sample processed

3 mL sterile syringes (two for each sample processed)

18-gauge needles (one for each sample)

25-gauge needles (one for each sample)

RNA Isolation Kit (Thermo Fischer Scientific, 12183025)

DNAse/RNAse-free water

70% ethanol made in DNAse/RNAse-free water

1.5 mL microcentrifuge tube

GeneAmp® 10X PCR Buffer II & MgCl₂ (Thermo Fischer Scientific, 1710071)

Random hexamers (Thermo Fischer Scientific, 2071896)

10 mM dNTP Mix (Thermo Fischer Scientific, 2079395)

Multiscribe™ Reverse Transcriptase (Thermo Fischer Scientific, 00751040)

RNase inhibitor (Thermo Fischer Scientific, 2032166)

2X PCR SuperMix (Quantabio, #95136)

Primers (100 μM, see Support Protocol 3)

PCR tubes (.2 mL)

Agarose

1X TAE buffer (see recipe in Reagents and Solutions)

SYBER® Green (Bio-Rad, 172–5120)

Primers (10 μM, see Support Protocol 3)

Vortex mixer

Microcentrifuge

Spectrophotometer capable of providing A260/A280 and A260/A230 ratios

Mini-centrifuge (able to fit PCR tubes)

PCR machine

Microwave

Electrophoresis power supply (e.g. PowerPac™ Universal Power Supply, Bio-Rad)

Submerged horizontal electrophoresis cell (e.g. Wide Mini-Sub Cell GT Cell, Bio-Rad)

Agarose gel casting kit (e.g. Wide Mini Handcasting Kit, Bio-Rad)

Gel Imaging System

96-well PCR plate and seal

Centrifuge with temperature control and ability to fit a 96-well PCR plate

Real-time PCR machine

Protocol 5.1 – RNA Isolation

• 1Clean gloves, working surface, and pellet pestles with DNAse/RNAse removal solution and wipe with a delicate task wipe. Then wipe down with 70% ethanol in ultrapure water. Cover pellet pestles with a delicate task wipe until ready for use.
• 2Follow the steps listed in the PureLink™ RNA Mini Kit manual to isolate the RNA.
• 3
  Determine the quality of each RNA sample using a spectrophotometer. Ideal values are listed below:

  1. A260/280 ≥ 1.8
  2. A260/230 ≥ 1.5

     While these values are ideal, RNA of lesser quality can be used if a control gene is used in addition to the target gene. RT-PCR and qRT-PCR results from the control gene can be used to determine if a particular sample is too poor to use.

• 4Store RNA at −80°C or keep on ice and proceed to protocol for cDNA synthesis.

Protocol 5.2 – cDNA Synthesis

• 5Thaw the 10X PCR Buffer (no MgCl₂), MgCl₂, random hexamers, and dNTPS on ice. (The RNAse inhibitor and reverse transcriptase should remain at −20°C until immediately before use).
• 6While the reagents are thawing dilute each RNA sample to 20 ng/μL. Test the quality and record the diluted concentration for each sample. The concentrations for all samples being run together should be within 5 ng/μL of each other for more accurate comparisons.
• 7Once the reagents are thawed, prepare the cDNA master mix (see recipe in Reagents and Solutions).
• 8Aliquot 20 μL of the cDNA master mix into labeled PCR tubes (one for each sample).
• 9Add 5 μL of the respective diluted RNA sample in to each PCR tube.
• 10Gently centrifuge PCR tubes for ~1 s to ensure all liquid is at the bottom of the tubes.
• 11
  Run the cDNA synthesis PCR protocol (the following is based on the cDNA kit used in this protocol. Please see the instruction manual if not using the same reagents listed here):

  1. 25°C for 10 min
  2. 48°C for 30 min
  3. 95°C for 5 min
  4. 4°C for ∞
• 12Store cDNA at −20°C or proceed to RT-PCR or qRT-PCR.

Protocol 5.3 – RT-PCR and Gel Electrophoresis

• 13
  Follow steps as listed under Protocol 4.2 and 4.3 of Basic Protocol #4. Note that for this RT-PCR, the primers used are different from those used previously.

  1. See Support Protocol 3 for steps on designing these primers

Protocol 5.4 – qRT-PCR

• 14Thaw SYBR® Green and primers on ice.
• 15While the reagents are thawing, set the centrifuge to 4°C and allow to cool to temperature.
• 16As the centrifuge cools, make a master mix for each primer set (see recipe in Reagents and Solutions).
• 17
  Aliquot 19 μL of each master mix into individual wells in the 96-well plate. Visually check each well to ensure equal volume across all wells.

  To improve the reliability of results, reactions should be done in triplicate. Thus, there should be three wells per sample per master mix. If there are two primer sets – target gene and control gene – there should be six wells per sample.

• 18Add 1 μL of cDNA to the 19 μL master mix aliquots and pipette to mix.
• 19Place seal over plate. Ensure the seal fully adheres to plate by gently going over each well with delicate task wipe.
• 20Centrifuge plate at 4°C at 2,500 rpm for 5 min.
• 21
  Place plate in real-time PCR machine and run PCR protocol (the following is based on the reagents, primers, and machine used in this protocol. Please see the instruction manual if not using the same items listed here):

  1. 95°C for 5 min
  2. 40 cycles of 95°C for 15s and 60°C for 30s
  3. Default melt curve (4°C ∞)
• 22Once complete, use the C_T values obtained to calculate relative expression using the ΔΔC_T method (Livak & Schmittgen, 2001).

Support Protocol 3: Designing primers for RT-PCR and qRT-PCR

Validation using RT-PCR or qRT-PCR requires detecting the presence of the targeted gene and a control gene. Only two primer sets are required for these PCRs. The first set will detect the target gene; the second will detect the control gene. Each primer should be ~15–20 bps, and the resulting PCR product for both reactions should be ~500 bps or less. Once the primer sequences are determined, primers can be ordered online.

Primer Set 1: Detecting the target gene.

The forward and reverse primers should begin just before and after, respectively, a sequence within the target gene. PCR product should be present for control tissue and absent for knockout tissue.

Primer Set 2: Detecting the control gene.

The forward and reverse primers should begin just before and after, respectively, a sequence within the control gene. There should be PCR product for all tissue.

BASIC PROTOCOL 6: VALIDIATION OF TISSUE-SPECIFIC KNOCKOUT USING WESTERN BLOT ANALYSIS

In addition to validating tissue-specific knockouts at the genomic and mRNA level, it is also beneficial to validate knockouts at the protein level. If an antibody for the targeted protein is available, protein levels in animal tissue can be assessed using Western blot analyses. If a knockout was successful, there should be decreased protein levels in the target tissue of knockouts compared to controls; while protein levels in non-target tissue of knockouts should be comparable to controls.

Western blot analysis involves isolating protein from a tissue samples, measuring the protein concentration, separating proteins using gel electrophoresis, transferring those proteins to a nitrocellulose membrane, and probing a specific protein using antibodies. The following protocol is a general protocol for Western blotting developed within the Mukhopadhyay Lab.

Tissue collection should be performed with RNAse- and DNAse-free materials. Once extracted, place tissue in a 1.5 mL microcentrifuge tube. Record the wet weight of the tissue. Tissue can be immediately processed for protein isolation (detailed below) or flash frozen in liquid nitrogen and stored at −80°C.

Materials

Fresh or fresh frozen tissue

1X lysis buffer (see recipe in Reagents and Solutions)

Protease inhibitor mini tablet (Thermo Scientific, A32953)

Protein Assay Dye Reagent Concentrate (Bio-Rad, 5000006)

2 mg/mL protein standard

Deionized water

30% degassed Acrylamide/Bis

1.5 M Tris(hydroxymethyl)aminomethane hydrochloride (Tris-HCl), pH = 8.8

0.5 M Tris-HCl, pH = 6.8

10% w/v Sodium dodecyl sulfate (SDS)

10% Ammonium persulfate (APS)

TEMED

70% ethanol

6X Sample Buffer (see recipe in Reagents and Solutions)

2X Sample Buffer (see recipe in Reagents and Solutions)

SDS-Page Buffer (see recipe in Reagents and Solutions)

Protein ladder

Transfer Buffer (see recipe in Reagents and Solutions)

Ponceau S

1X Tris-Buffered Saline + Tween (TBST) (see recipe in Reagents and Solutions)

Non-fat dry milk (Bio-Rad, 1706404)

Primary antibody to probe target protein

Secondary antibody to probe primary antibody

Detection Reagent 1 (Thermo Scientific, 1859701)

Detection Reagent 2 (Thermo Scientific, 1859698)

1.5 mL microcentrifuge tubes

Disposable spectrophotometer cuvettes

Gel-loading pipette tips

Glass dish (~9 X 9 or larger)

Small spatula

Small roller

Large ice bucket (able to fit the electrophoresis transfer cell)

Plastic square petri dish (Electron Microscopy Sciences, 70691)

Sonicator

Microcentrifuge

Spectrophotometer with ability to read absorbance of 595 nm

Heat block with water bowl

Mini-PROTEAN® Tetra Cell (Bio-Rad, 1658003)

Electrophoresis Power Supply (e.g. PowerPac™ Universal Power Supply, Bio-Rad)

Mini Trans-Blot® Electrophoretic Transfer Cell (Bio-Rad, 1703930)

Table rocker

Gel Imaging System (GE Amersham Imager 600QC)

Protocol 6.1 – Protein Isolation

• 1Prepare 10 mL of 1X lysis buffer in PBS from the 10X lysis buffer (see recipe in Reagents and Solutions) and place on ice. Dissolve one protease inhibitor mini tablet in the 1X lysis buffer.
• 2
  Place tissue sample on ice. Add 100 μL of aliquoted 1X lysis buffer (containing protease inhibitor) for every 1 mg of tissue.

  If the wet weight of the tissue is more than 10 mg, transfer the tissue to a larger tube before adding lysis buffer. Alternatively, cut the tissue, cut the tissue and use the new wet weight to determine the amount of lysis buffer needed.

• 3Homogenize sample by sonicating at 40% output twice for 1 min with a 1 min interval between. Keep sample on ice. The sample should not get warm.
• 4Centrifuge at 10,000 rpm for 10 min at 4°C.
• 5Collect the supernatant and pipette into a new 1.5 mL tube (or larger tube if necessary). Place on ice and proceed to measuring the protein concentration.

Protocol 6.2 – Measuring protein concentration (Bradford assay)

• 6Dilute the Bio-Rad Protein Assay Dye Reagent Concentrate to 1X. Vortex to mix.
• 7Label five 1.5 mL microcentrifuge tubes: “0 μL”, “1 μL”, “2 μL”, “4 μL”, “8μL”. Label additional 1.5 microcentrifuge tubes to designate tissue samples.
• 8Aliquot 1 mL of the 1X Protein Assay Dye into each microcentrifuge tube.
• 9
  Add the respective volume of the 2 mg/mL protein standard to the aliquoted Protein Assay Dye (i.e. for the tube labeled “0 μL”, add 0 μL of protein standard; for the tube labeled “1 μL”, add 1 μL of protein standard). Add 1 μL of tissue sample supernatant into respective tubes. Vortex all to mix.

  When using a 2 mg/mL protein standard, these volumes provide 0 μg, 2 μg, 4 μg, 8 μg, and 16 μg, respectively. Alter volumes as necessary to ensure sufficient protein is added to the dye if a different concentration of protein standard is used.

• 10Transfer mixtures to individual cuvettes.
• 11
  Measure absorbance at 595 nm for each using the spectrophotometer. Record each value.

  Use the “0 μL” to blank.

• 12
  Calculate the concentration of protein for each tissue sample.

  1. Calculate a linear equation (y = mx + b) using the values from the protein standard, where x is μg protein and y is absorbance.
  2. Ensure the calculated slope is between .96 and 1. This is an indicator of accuracy and whether results from the Bradford assay can be trusted.
  3. For each tissue sample, use the respective absorbance value as y, and solve for x. The protein concentration is x μg/1 μL.

Protocol 6.3 – SDS-Page

• 13
  Cast a 10 % resolving gel with a 5% stacking gel by completing the following steps:

  The gel percentage (percent of acrylamide) required depends on the sizes of the proteins being separated. Alter the amount of acrylamide and deionized water if a different gel percentage is required.

  1. Assemble the gel casting apparatus using a gel cast for 1.0 mm well size.

     If needed, ensure the apparatus is properly sealed by filling the gel cast with water. Wait approximately 15 min. If no water has leaked out, pour out the water, using a delicate task wipe to absorb any remaining water. Proceed to the next step. If water has leaked out, reassemble the apparatus and test with water again.

  2. Mix together the following for the resolving and stacking gels:

     1. 10% Resolving Gel: 2.05 mL deionized water, 1.65 mL degassed Acrylamide/Bis, 1.25 mL 1.5 M Tris-HCl (pH = 8.8), 50 μL 10% SDS
     2. 5 % Stacking Gel: 2.85 mL deionized water, 850 μL degassed Acrylamide/Bis, 1.25 mL 0.5 M Tris-HCl (pH = 6.8), 50 μL 10% SDS
  3. Add 50 μL of 10% APS and 5 μL of TEMED to the 10% resolving gel mixture. Pipette to mix. Then, using a P1000 pipette and tip, slowly add the 10% resolving gel mixture to the gel cast. Fill up to approximately 1 in from the top, so there is ample room for the stacking gel and wells. Keep any remaining gel mixture.
  4. Add 70% ethanol to the gel cast (this will help level the resolving gel).
  5. Wait approximately 20 min for the resolving gel to harden.

     Use any remaining gel liquid to check if the resolving gel has solidified.

  6. Pour out the 70% ethanol.
  7. Add 50 μL of 10% APS and 5 μL of TEMED to the 4% stacking gel mixture. Pipette to mix. Then, using a P1000 pipette and tip, slowly add the 4% stacking gel mixture to the gel cast. Fill to the top of the gel cast. Be careful not to introduce any bubbles into the gel. Keep the remaining gel mixture.
  8. Add a 1.0 mm well comb.
  9. Wait approximately 20 min for the stacking gel to harden.

     1. Use any remaining gel liquid to check if the resolving gel has solidified.
• 14While waiting for the gel to harden, fill a water bowl with water and place on heat block. Set to maximum temperature, and wait for the water to boil.
• 15As the water is heating, prepare tissue samples, by mixing 10 μL of 2X sample buffer with 10 μL of protein sample. Boil for ~10 min.
• 16Once the gel has hardened, remove the gel cast plates from the gel casting stand and frames. Do not remove casting plates from gel.
• 17
  Secure casting plates (with gel) in the electrode assembly. If only one gel is being used, place the buffer dam on the other side of the electrode assembly. Place in the electrophoresis cell tank. Be sure to align black to black and red to red.

  To ensure that the casting plates and electrode assembly are properly sealed, fill the space between the casting plates (or casting plate and buffer dam) with SDS page buffer (see recipe in Reagents and Solutions). Allow to sit for a few minutes. If buffer leaks into the tank, the assembly is not well sealed. Pour our remaining SDS Page Buffer, unclip the casting plate(s) and reassemble. Place back in tank, fill with SDS page buffer, and check for leaking. Repeat until a proper seal is achieved.

• 18Fill the tank, up to the max line, with SDS Page Buffer.
• 19Carefully remove wells from the gel plates.
• 20Load ~5 μL of protein ladder into well using a gel-loading pipette tip.
• 21Load the necessary volume of boiled sample(s) (~30–50 μg of protein) into wells using gel-loading pipette tips.
• 22Place top on the electrophoresis cell tank, aligning black to black and red to red. Insert electrodes into the electrophoresis power supply and run gel at ~160 V for 60 min, or until the dye front reaches the bottom of the gel. Do not let the dye front run off of the gel.

Protocol 6.4 – Transfer

• 23Fill the ice bucket with ice, leaving room for the electrophoresis cell tank to be surrounded by ice.
• 24Pour ~500 mL of Transfer Buffer (see recipe in Reagents and Solutions) into the glass dish
• 25
  Disassemble the SDS Page electrode assembly. Carefully remove the short plate from the gel.

  Use a small spatula to gently lift the top of the short plate. This will lightly loosen the short plate from the gel making it easier to remove.

• 26Cut of the stacking gel and place gel (with spacer plate) into glass dish with transfer buffer.
• 27
  Remove the gel from the space plate and float in the transfer buffer.

  Use the spatula to gently lift the top of the gel from the spacer plate. Then gently remove spacer plate. Be careful not to rip the gel.

• 28Soak two foam pads and two pieces of filter paper in Transfer Buffer.
• 29
  Assemble transfer apparatus

  1. Place the color-coded cassette into the glass dish with transfer buffer, with the clear side against the bottom of the dish. Be careful to avoid the gel, so it does not get damaged.
  2. Place one pre-soaked foam pad on top of the clear side of the cassette.
  3. Place one piece of pre-soaked filter on top of the foam pad.
  4. Soak the nitrocellulose membrane in transfer buffer and place on top of the filter paper. Pour some transfer buffer over the membrane to remove any bubbles.
  5. Gently place the gel on top of the nitrocellulose membrane.
  6. Place the second pre-soaked filter paper on top of the gel.
  7. Place the second pre-soaked foam pad on top of the filter paper.
  8. Using a small roller, roll over the foam pad once from left to right to remove any bubbles.
  9. Place the black side of the cassette down and clamp the transfer apparatus using the white clip.
• 30Place transfer apparatus into the electrode module, with the black sides facing each other. Place the electrode module with transfer apparatus into the electrophoresis cell tank and place a frozen ice pack into the electrophoresis cell tank.
• 31Fill tank with remaining Transfer Buffer, and place the entire apparatus into the ice bucket.
• 32Place the top on the electrophoresis cell bucket, aligning black to black and red to red. Run transfer at 100 V for 60 min.

Protocol 6.5 – Immunoblotting

• 33Disassemble transfer apparatus. Place the nitrocellulose membrane face up in a square plastic petri dish.
• 34Pour Ponceau S over the membrane, place dish on a rocker, and stain for ~2 min. Pour out Ponceau S and wash membrane with water until protein bands are visible to ensure successful protein transfer. Wash any remaining Ponceau S with 1X TBST.
• 35Prepare 100 mL of a 5% (w/v) non-fat dry milk solution in 1X TBST and 100 mL of a 1% (w/v) non-fat dry milk solution in 1X TBST.
• 36Pour out 1X TBST covering membrane, and cover membrane in the 5% dry milk solution. Sit at room temperature on a rocker for 60 min to block any non-specific binding.
• 37Approximately 5 min before the membrane finishes blocking in the 5% dry milk solution, prepare a 1:1000 dilution of the primary antibody in 10 mL of the 1% dry milk solution (alter the dilution ratio if needed). Store remaining 1% dry milk solution at 4°C.
• 38Pour out 5% dry milk solution, and cover membrane in diluted antibody solution. Incubate overnight at 4°C on a rocker.
• 39Pour out diluted antibody solution, and wash membrane in 1X TBST for 60 min on a rocker at room temperature, changing to new 1X TBST ~every 12 min (5 times).
• 40Prepare a 1:3000 dilution of the secondary antibody in 10 mL of the 1% dry milk solution (alter dilution ratio if necessary).
• 41Pour out 1X TBST and cover membrane in the diluted secondary antibody solution. Incubate for 40 min at room temperature on a rocker.
• 42Pour out diluted secondary antibody solution and wash membrane in 1X TBST for 60 min on a rocker at room temperature, changing to new 1X TBST ~every 12 min (5 times).
• 43Pour out 1X TBST and cover membrane in a 1:1 solution of Detection Reagent 1 and Detection Reagent 2 (~ 4 mL).
• 44Image on the gel imaging system using the chemiluminescense setting.

REAGENTS AND SOLUTIONS

cDNA Master Mix

For 1 reaction, combine the following and pipette to mix (do not vortex):

• 4.5 μL 10X PCR Buffer

• 6.5 μL MgCl₂

• 1.25 μL dNTPs

• 1.25 μL Random hexamers

• 7.35 μL RNAse-/DNAse-free ultrapure water

• 0.50 μL RNAse inhibitor

• 0.65 μL Reverse Transcriptase

Add the RNAse inhibitor and reverse transcriptase last, and do not remove from the freezer until immediately before adding to the master mix.

Make enough of the master mix to complete all necessary PCR reactions and still have some master mix remaining. Keep master mix on ice during use.

ES Cell Medium

500 mL Knockout DMEM

91.5 mL FBS

6.1 mL 1X Glutamine

6.1 mL 1X Pen-Strep

6.1 ml 1X BME

Store at 4° C.

ES Cell Injection Medium

88 mL Knockout DMEM

10 mL FBS

2 mL HEPES

Store at 4° C.

2X Freexing Medium

12 mL Knockout DMEM

4 mL FBS

4 mL Dimethyl Sulphoxide (Sigma D2650)

Make fresh.

5X Gel Buffer

151 g Tris Base

720 g Glycine

Fill up to 10 L with MilliQ or deionized water.

Store up to six months at room temperature.

Hybridization Buffer

25 mL 20X SSC

10 mL10% SDS

2 mL 1M NaPO4, pH 7.0

2 mL 5 mg/mL herring sperm DNA

500 mg dry milk

50 mL formamide

11 mL sterile water

Make fresh.

LongAmp PCR Cocktail mix

5 μL LongAmp Taq 2X Master Mix

0.5 μL 10 μM cassette primer

0.5 μl 10 μM genomic primer

3 μL water

Make a master mix of the components sufficient for the number of clones to be screened plus 10%.

Lysis Buffer (Protocol 1.3)

For each plate:

• 0.05 mL 1M Tris, pH 7.5

• 0.01 mL 5M NaCl

• 0.1 mL 0.5 M EDTA

• 0.25 mL 10% Sarkosyl

• 0.5 mL 10 mg/mL Proteinase K

• 4.09 mL sterile water

• Make Fresh.

10X Lysis Buffer (Protocol 6.1)

• 20 mL 1M Tris (pH = 7.4)

• 10 mL Triton X-100

• 10 mL 10% (w/v) SDS

• Fill up to 100 mL with 1X PBS (see above recipe)

• Store up to one year at 4°C.

PCR Master Mix

For 1 PCR reaction, combine the following and vortex to mix:

• 10 μL AccuStart II PCR SuperMix (2X)

• 10 μL RNAse-/DNAse-free ultrapure water

• 0.2 μL 100 μM Forward primer

• 0.2 μL 100 μM Reverse primer

Make enough of the PCR Master Mix to complete all necessary PCR reactions and still have some master mix remaining. Keep master mix on ice during use. Use this recipe for both Protocol 4.2 and Protocol 5.3.

1X Phosphate-Buffered Saline (PBS)

Make a 10X PBS stock solution by mixing the following:

• 80 g NaCl

• 2 g KCl

• 9.16 g Na₂HPO₄

• 2 g K₂HPO₄

• Fill up to 1L with MilliQ water.

To make 1L of 1X PBS, dilute 100 mL of 10X PBS in 900 mL of MilliQ water, and pH to 7.3.

Both 10X and 1X PBS can be stored up to six months at room temperature.

qRT-PCR Master Mix

For 1 reaction, combine the following and vortex to mix:

• 10 μL SYBR® Green

• 8.2 μL RNAse-/DNAse-free ultrapure water

• 0.40 μL 10 μM Forward Primer

• 0.40 μL 10 μM Reverse Primer

Make enough of the master mix to complete all necessary PCR reactions and still have some master mix remaining. Keep master mix on ice during use.

6X Sample Buffer

25 mL Glycerol

5 g SDS

7.8 mL 1 M Tris (pH = 6.8)

25 mg Bromophenol blue

Store at room temperature.

2X Sample Buffer

500 μL 6X Sample Buffer (see above recipe)

1 mL 8 M Urea

100 μL 0.25 M Dithiothreitol (DTT)

75 μL BME

Store at room temperature for short-term use (1 week) or store at −20°C for long-term storage.

SDS-Page Buffer

200 mL 5X Gel Buffer (see above recipe)

10 mL 10% (w/v) SDS

Fill up to 1 L with MilliQ water

Store at room temperature.

STO Medium

500 mL Knockout DMEM

38.5 mL FBS

5.5 mL Glutamine

5.5 mL Pen-Strep

1X TAE Buffer

Make a 50X TAE Buffer stock by mixing the following:

• 242 g Tris Base

• 57.1 mL Glacial Acetic Acid

• 100 mL 0.5 M EDTA (pH = 8.0)

• Fill up to 1 L with MilliQ water

To make 1L of 1X TAE Buffer, mix 20 mL of 50X TAE with 980 mL of MilliQ water.

Store at both the 50X and 1X TAE Buffer for up to six months at room temperature.

1X TBST

8.7 g NaCl

10 mL 1M Tris (pH = 7.4)

0.5 mL Tween 20

Fill up to 1 L with MilliQ water.

Store up to six months at room temperature.

Transfer Buffer

200 mL 5X Gel Buffer (see above recipe)

200 mL Methanol

Fill up to 1 L with MilliQ or deionized water.

Store up to six months at room temperature.

COMMENTARY

Background Information

Developed in the 1980s and 1990s, the Cre-loxP system has become indispensable for the generation of tissue-specific knockout mice. The Cre enzyme and loxP do not occur naturally in mammalian genomes, and were initially derived from bacteriophage (Klos, 2004; Sauer, 1998). In the late 1980s, Dr. Brian Sauer first introduced the Cre-loxP system to yeast, demonstrating site-specific recombination in a eukaryotic cell (Sauer, 1987). Dr. Sauer, along with Dr. Nancy Henderson, then introduced the Cre-loxP system to a mouse cell line, demonstrating functional site-specific recombination in mammalian cells (Sauer & Henderson, 1988). These studies provided initial evidence that the bacterial-derived Cre-loxP system could be used to target specific genes in a variety of cell types in vitro, providing a basis for testing its function in while organisms.

One of the first tissue-specific knockout mice was generated in the early 1990s by Dr. Jamey Marth. Dr. Marth showed that the Cre enzyme could be inserted in the mouse genome under the control of a tissue-specific promoter, the thymus promoter lck, and that loxP sites could be integrated to flank a specific gene, β-galactosidase. The transgenic mice expressing both Cre and the floxed gene exhibited loss of β-galactosidase in thymus cells (Orban, Chui, & Marth, 1992). The successful generation of a tissue-specific knockout mouse using the Cre-loxP system provides a novel and revolutionary method for modeling human disease and studying the tissue-specific functions of proteins.

In addition to gene targeting in mouse ES cells as in the Cre-loxP system, it is also possible to generate mice with conditional alleles using CRISPR-Cas9 (Yang et al., 2013). Two benefits of using CRISPR-Cas9 are that it is less labor intensive to make potential founder mice, compared to gene targeting where extensive ES cell culture and screening work is required at the start; and possible founder animals can be screened as early as four to six weeks after microinjection. However, CRISPR-Cas9 does have limitations. The efficiency of obtaining a founder mouse with two independent loxP sites integrated correctly on the same chromosome can vary (our unpublished data and personal communications from other transgenic core labs). Low efficiencies are likely due in part to non-homologous end joining (NHEJ) repair being more favored over homology-directed repair following double-strand DNA cleavage by Cas9 (Singh, Schimenti, & Bolcun-Filas, 2015). A second difficulty with the CRISPR approach is that once potential founders are born, detailed PCR and sequence analysis is required along with time-consuming breeding steps. Founder animals are usually mosaic and can transmit multiple alleles (Yen et al., 2014). Thus, founder animals need to be extensively characterized by PCR and sequence analysis to identify animals that contain a correct floxed allele and no other alterations (Kosicki et al., 2018). Lastly, there is also the possibility of off-target mutations in CRISPR mice, although off-target mutations appear to be rare (Iyer et al., 2015). Overall, the CRISPR-Cas9 system provides an alternative method for creating conditional alleles, but efficiencies can vary and extensive validation is required to ensure only the intended alterations have occurred.

Critical Parameters

In order to validate tissue-specific knockouts using RT-PCR or qRT-PCR, preserving the quality of RNA analyzed in these produces is vital. RNA is easily and quickly degraded, and poor quality or quantity of RNA decreases the reliability of RT-PCR and qRT-PCR. All working surfaces and materials must be cleaned with a DNAse/RNAse removal solution. Isolated RNA must also be kept on ice while in use and at −80°C for long-term storage to slow down degradation. If results from the RT-PCR or qRT-PCR are inconsistent, measure the RNA concentration and quality again to ensure excessive degradation has not occurred.

For both the PCR and RT-PCR, if the band(s) for the PCR product are not readily detectable, troubleshoot by altering the PCR conditions. If optimizing the PCR conditions does not work, re-check the PCR primers and design new primers if needed.

When conducting qRT-PCR, accurate pipetting is necessary to ensure reliable results. Because only 1 μL of cDNA is used for each reaction, it is critical that each reaction has exactly 20 μL of solution (1 μL cDNA and 19 μL of master mix). If qRT-PCR results are variable but the RNA quality is not a concern, assess accuracy and precision of the experimenter’s pipetting.

For Western blotting, it is important to remember that the SDS-page gel is delicate. Precautions must be taken to ensure the gel does not rip during the procedure. Submerging the gel in liquid during removal and transfer aid with this.

Statistical Analyses

Statistical analysis is needed to validate results obtained in qRT-PCR. Depending on the total number of groups being analyzed, varying statistical tests may be appropriate. For this protocol, relative expression from only two groups are compared: control and knockout animals. Thus, a t-test is appropriate for analyzing these results.

Understanding Results

For gene targeting, the number of targeted clones obtained will vary depending on the gene and the targeting vector, but efficiencies of 5–20% are typical. The percentage of clones that contribute to the germline in chimeric mice can also vary, but our experience is that more than half of the hybrid or 129 ES clones go germline and approximately 40% of the C57BL/6N KOMP/EUCOMM clones go germline. Given this frequency, it is recommended to purchase at least three KOMP or EUCOMM clones for your gene of interest if they are available.

The efficiency of the IVF will depend on the quality of the cryopreserved sperm. We ask the providing repository for information on the fertilization rate of the sperm as this information helps us plan for how many mice to use for oocyte recovery. For C57BL/6 mice, we estimate to recover approximately 20 oocytes per superovulated mouse. For sperm with a reported fertilization rate of 50%, we would plan to superovulate six mice to get about 120 oocytes. With a predicted fertilization rate of 50%, there should be approximately 60 two-cell embryos the following day after IVF. The embryos are then transferred to three or four 0.5 day pseudopregnant female mice. About 30 mice should be recovered from the transfers.

The results obtained for validating tissue-specific knockouts rely on the presence or absence of PCR products or a protein bands. Results should indicate mice designated as knockouts (Basic Protocol 3) exhibit loss of the target gene only in the tissue associated with the promoter controlling the Cre expression (Basic Protocols 4 – 6). An example of results obtained from Basic Protocols 4 – 5 can be found in Figure 2 A-C of Taylor et al. 2019.

Time Considerations

It takes approximately nine to twelve months to generate a mouse with a floxed allele by gene targeting in ES cells if everything goes as planned. The IVF procedure takes one week to complete and 19 days until pups are born after embryo transfer. Recovered Cre transgenic lines are ready for crossing to floxed allele mice in 8–10 weeks. Once that litter is born and is ready for breeding (~3 months), heterozygous mice are then crossed with floxed mice to produce the knockout strain (~3 weeks).

Validation of the generated knockouts takes considerably less time. A list of the time expected to complete each protocol are as follows: Validation using genomic DNA – 4 hours; RNA extraction – 1 hour; cDNA synthesis – 2.5 hours; RT-PCR – 2 hours; qRT-PCR – 3 hours; Western Blot – 6 hours.

Significance Statement.

Tissue-specific knockout mice are an essential tool in toxicology research. The ability to remove gene function from a particular tissue, while the leaving function intact in other tissues, can lead to unexpected findings on the role of specific tissues in the development and progression of disease pathology. This protocol gives a detailed description of how to generate and validate tissue-specific knockouts.