Giancarlo Ghiselli and Renato V Iozzo

1. Introduction

1.1. Overview of Gene Targeting in Mammalian Cells

Gene targeting allows the generation of specifically designed mutations in cells by the mean of homologous recombination between exogenous DNA and endogenous genomic sequences (1-3). This is possible because a fragment of genomic DNA introduced into a cell can locate and recombine with the chromosomal homologous sequences (4). Although the mechanisms of exogenous DNA transfer to the nucleus and of homologous recombination are still poorly understood, important advances have been made in identifying the requirements for the targeting vector and the desirable features of the targeted locus that maximize gene targeting (5). Presently, various gene targeting techniques can be successfully applied to a variety of organisms. In mammals, gene targeting strategies have been developed mainly for the genetic manipulation of mouse embryonic stem (ES) cells. Mating of the transgenes leads to the generation of animals in which the functional significance of a specific gene silencing can be investigated in different conditions. Cell lines can also be established from mouse organ biopsies. There are, however, important limitations to the attainability of cell lines of interest from transgenic animals. First, ES gene knockout techniques are currently limited to the mouse. Because there are major interspecies differences with regard to gene pattern of expression and implication in disease, extrapolation from the results obtained with mice cells to the pathophysiological mechanisms in humans is not warranted. This is particularly the case when genes involved in malignant transformation and development are investigated. Second, the availability of established cell lines bearing specific mutations makes somatic gene targeting highly desirable. In this case the implication of the expression of a particular gene on the cell behavior can be assessed directly against a well-defined genetic background. Third, the establishment of an organ-specific cell line involves complex genetic rearrangements leading to immortalization. In other words, cell lines, even if

From: Methods in Molecular Biology, Vol. 171:Proteoglycan Protocols Edited by: R. V. Iozzo © Humana Press Inc., Totowa, NJ

derived from the same animal cannot be considered isogenic in the sense that the pattern of expression of several genes — as opposed to the single targeted gene—is affected. In addition, the genetic mutation that has been introduced through homologous recombination in ES cells may be lethal even at a stage preceding embryonic development, thus precluding the possibility of establishing a cell line. Finally, the cost of originating and maintaining a transgenic animal colony is high.

These potential drawbacks are particularly relevant when considering the gene knockout of proteoglycan core proteins. There is in fact ample evidence that proteoglycans play a crucial role, both in development as well as in modulating the phenotypic expression of somatic cells. This is a case where the ES cell knockout strategy is of dubious value, as the generated transgenic animals will likely display a phenotype that is the result of a constellation of effects ensuing at different stages of life. A mechanistic interpretation of the function of a gene at a cellular level is thus difficult. This is more so in the case of diseases in which an array of genes rather than a single gene play a role, such as in cancer.

The subject of homologous recombination and gene targeting has been reviewed extensively in the recent past (5,6). The reader is directed to that literature for a critical examination of the various strategies devised for achieving successful gene targeting and knockout. In this chapter, we will discuss those aspects of the targeting vector design that are pertinent to somatic gene knockout and the methodology for vector transfer into established cell lines other than ES cells. The design of an effective screening strategy for the identification of homologous recombinants by genotyping and functional assays is also discussed. As proteoglycans play a role in cell differentiation and growth (7) their gene knockout can significantly affect cell behavior. Consequently, care should be taken in devising a screening strategy that is not negatively biased toward the selection of clones with a rate of growth different from that of the parent cell line, as this would lead to poor recovery of the clones in which homologous recombination has occurred.

1.2. Targeting Strategies for Gene Knockout

A major limitation of gene targeting is that DNA introduced into the mammalian genome integrates by random, nonhomologous recombination at a rate 100 to 100,000 times more frequently than by homologous recombination. In order to identify and recover the small fraction of cells in which a homologous recombination occurs, several strategies have been developed. Through these strategies, successful homologous recombination have been reported to occur in at about 1% of stable transfected ES clones. Two of these strategies, the so-called positive-negative selection (PNS) (8) and the promotorless strategy (9,10) have been found to be particularly advantageous. The PNS approach exploits the use of two selectable markers in the targeting vector. The first marker gene harboring a drug resistance cassette is inserted within the region of homology of the vector and is driven by an exogenous promoter. This marker serves to disrupt the gene translation and at the same time to select the cells that have incorporated the homologous DNA region of the targeting vector within the chromosomal DNA. Examples of selectable markers are neo, which confer resistance to G418, an analog of neomycin that is toxic to mammalian cells, and hydro and puro, which confer resistance to the hygromycin and the puromycin antibiotic, respectively. The second marker's gene, also driven by an exogenous promoter, is placed outside the homology region of the targeting vector and confers sensitivity to a toxic agent (e.g., Herpes simplex virus thymidine kinase [HSV-i£] which impart sensitivity to purine analog such as ganciclovir, FIAU, and acyclovir through phosphorylation of these agents). The rationale is that, unless a double reciprocal crossover (i.e., a replacement) is taking place and the HSV-i£ gene is excised out of the inserted vector, cells become sensitive to the purine toxic agent as the negative-selection gene is incorporated into the chromosomal DNA. The main advantage of the PNS vector strategy is that there is no requirement for the targeted gene to be expressed, as both markers gene are driven by their own promoters. This strategy has been adopted extensively in ES cells. The occurrence of homologous recombination is somatic cell lines is, however, two to three orders of magnitude lower than in ES cells (5) and targeting strategies have been developed that allow the identification of homologous recombinant events with higher efficiency than that allowed by the PNS strategy. The attainment of high screening efficiency is crucial for somatic gene knockout inasmuch as a double round of gene targeting is required. The so-called promotorless selectable marker strategy relies on a single selectable marker that becomes activated if the targeting vector is incorporated in a chromosomal region that allows the gene to be driven by an endogenous promoter. The expected transcriptional product is represented by a fusion protein comprising the N-terminal region of the targeted protein and the selectable marker. Since activation by cellular promoters occurs in about 1% of random integration events, the use of promotorless vectors provides a 100-fold enrichment for homologous recombination by reducing the background of nonhomologous recombination. By careful manipulation of the concentration of the selection agent it has been possible to increase the rate of legitimate recombination to values approaching or even exceeding 10% (11). A recent survey of the literature (12) of 23 experiments of gene targeting with a promotorless vector has revealed that gene targeting frequency in human cell lines is highly variable, ranging from 1/3 for the p21 gene targeted in HCT116 colon carcinoma cells to 1/940 for the p53 gene in the same cell line, suggesting important locus-to-locus variability. A significant advantage of the promotorless strategy with reference to somatic gene knockout is that since the end point of the experiment is represented by the suppression of a cell phenotypic trait, assays relying on the expression of the targeted gene product can be considered for screening purposes. This can greatly facilitate the screening of the stable transfected clones, bypassing the problem of identifying the modality of gene mutation through genomic characterization of the targeted locus.

1.3. Design of a Promotorless Targeting Vectors for Somatic Knockout of Proteoglycan Genes

Although the description of the method is general, this approach can be applied to the somatic knockout of any given proteoglycan gene. A targeting vector is designed to recombine with and mutate a specific chromosomal locus. The construction of a promotorless targeting vector involves the selection of a suitable genomic fragment harboring an exon into which a positive selection cassette can be inserted in frame. The positive selection marker serves two functions: (1) to isolate the rare transfected cells among which there are those that have properly integrated DNA, and (2) to silence the gene by mutation. The selected genomic fragment must be free of any functional promoter/enhancer elements and of repeating sequences that might increase the rate of detectable spurious recombinations. The desirable features of the homologous sequence to be incorporated in the targeting vector are:

1. The presence of regions coding for the N-terminus of the protein such that the expression of a proteins with functional activity is rigorously precluded.

2. The absence of repetitive sequences in order to reduce the chances of scattered illegitimate recombination in the chromosomal DNA (4).

3. The coding region harboring the marker should be located 5 kb or more from the transcrip-tional starting site. This is to prevent the gene promoter influencing the expression of selectable marker (5,11).

4. The selectable marker should be preferably inserted within an exon initiating and terminating in different splicing phase. This to avoid that splicing of the mutated exon might give rise to a functional gene product (13).

5. The size of the homologous sequence should be between 3 and 6 kb. Efficiency of homologous recombination is dependent on the length of the homologous region, but there is little evidence that beyond 6 kb the frequency of homologous recombination occurs at significantly higher rate (14-16). On the other hand, limitation in the size of the homologous region facilitates the construction of the vector

6. The rate of recombination is affected by the length of homologous end regions of the targeting vector (17). A nonmutated region of 500 bp at either end of the homologous region should provide sufficient linearized DNA area such that hybridization of exogeneous and chromosomal DNA at the site of crossover might occur efficiently.

7. The targeting vector needs to be linearized such that a single crossover event (insertion targeting) or rather a double crossover event (replacement targeting) is favored (18). Gene knockout by insertion occurs at a rate 10 times higher than by replacement (13). In order to favor insertion targeting, the targeting vector is linearized at a restriction site located within the region of homology. On the other hand, a replacement vector requires linearization outside the region of homology, within the vector backbone where unique cutting sites are already mapped. In this sense, the design of a replacement vector is facilitated compared to that of an insertion vector.

Detailed mapping of the region of homology is required, especially when the use of an insertion vector is contemplated. In-depth knowledge of the targeting site and its restriction fragment mapping is also required in order to generate DNA probes that can provide informative results on the successful integration of the targeting vector by Southern hybridization screening. Insertion and replacement targeting vectors give raise to different integration products (6). The genotype screening strategies should take into account that although some of the crossover events are favored over others, there is the possibility that a rather wide range of integration products is obtained, especially when using an insertion vector. This is because the pattern of integration is largely dependent on the DNA recombination and repair machinery of the host cell

Fig. 1. Integration pattern of the targeting vector into a genomic locus. A promotorless targeting vector is constructed by inserting a mutagen-selectable marker (illustrated by the heavy shaded box) into the coding region of the gene of interest (shown as lightly shaded boxes) such that a fusion protein terminated by the stop codon of the selectable marker is generated. The expression of the selectable marker (usually a gene whose product confer-resis-tance toward a cytotoxic drug) is driven by an endogenous cellular promoter coinciding with that of the targeted gene when a legitimate recombination takes place. Targeting vectors linearized at a site external to the region of homology integrate into the chromosomal DNA preferentially by replacing the endogenous DNA through a double crossover event (A). Gene disruption by insertion is favored when cells are transfected with a targeting vector that has been linearized within the region of homology (B). In this case the vector backbone becomes part of the mutated chromosomal locus. Because a single crossover event is necessary for the integration of foreign DNA, mutagenesis by insertion occurs at higher frequency than by replacement.

Fig. 1. Integration pattern of the targeting vector into a genomic locus. A promotorless targeting vector is constructed by inserting a mutagen-selectable marker (illustrated by the heavy shaded box) into the coding region of the gene of interest (shown as lightly shaded boxes) such that a fusion protein terminated by the stop codon of the selectable marker is generated. The expression of the selectable marker (usually a gene whose product confer-resis-tance toward a cytotoxic drug) is driven by an endogenous cellular promoter coinciding with that of the targeted gene when a legitimate recombination takes place. Targeting vectors linearized at a site external to the region of homology integrate into the chromosomal DNA preferentially by replacing the endogenous DNA through a double crossover event (A). Gene disruption by insertion is favored when cells are transfected with a targeting vector that has been linearized within the region of homology (B). In this case the vector backbone becomes part of the mutated chromosomal locus. Because a single crossover event is necessary for the integration of foreign DNA, mutagenesis by insertion occurs at higher frequency than by replacement.

line (4). Furthermore, there is clear evidence for a locus-to-locus susceptibility for integration of foreign DNA that affects both the rate of homologous integration as well as the pattern of integration (12). Generally, the final recovery product of a replacement vector is equivalent to the replacement of the homologous chromosomal sequence with all the components of the vector except for the regions flanking the homologous sequence, as all the heterologous part of the vector is excised and lost (17) (see Fig. 1A). The favored final product of an insertion vector on the other hand, is an increase in length of chromosomal DNA corresponding to the full length of the targeting vector including the vector backbone (see Fig. 1B).

1.4. Somatic Gene Knockout: Overall Strategy

After a suitable cell line and the targeting strategy have been selected, the somatic gene targeting involves the following general steps: (1) electroporation of the targeting vector into the host cell line, (2) selection of stable transfected cell colonies by drug selection, (3) rescue and growth of the cell colonies, (4) preparation of frozen stock and of duplicate plates for identification of the targeted clones by Southern blot analysis, PCR, or RT-PCR, (5) new round of targeting in the heterozygous clones utilizing a targeting vector harboring a new selectable marker, and (6) cloning, cell rescue, and gene analysis. The knockout is then confirmed by genotyping, Northern blot hybridization, and immunoassay techniques. For the construction of the promotorless targeting vector in p-Bluescript (Stratagene) or another suitable cloning vector, the neomycin cassette harboring the Neomycin phosphotransferase gene (neo) and its polyA tail is generated by PCR from a pSV2-Neo template using primers that allow for the in-frame ligation of the cassette into the selected coding region of the targeting vector. Following cloning and linearization of the construct, the targeting vector is introduced into the selected human cell line by electroporation using 50-100 ^g DNA per 107 cells. Human colon carcinoma HCT116 cells has been frequently used for this purpose (12). This cell line offers some key advantages over other normal or malignant cell lines. First, HCT116 carries a DNA mismatch repair-deficient gene, thereby enhancing the stability of the inserted foreign DNA (19). Second, these cells are sensitive to the cytotoxic effect of G418 over a relative wide range of concentrations (between 0.4 and up to 10 mg/mL), which is useful when attempting to improve the rate of recovery of successful transfectants by increasing the drug concentration (11). Third, unlike most transformed cell lines, HCT116 has a normal euploid, which implies that somatic knockout is completed in two rounds of gene targeting. Within 2 wk after electroporation and selection in G418, the drug-resistant colonies are ring-cloned and expanded. The targeting of the second allele is pursued by a second round of transfection with a targeting vector carrying a different drug resistance gene, usually hygro or puro. After genotyping of the rescued clones by Southern hybridization, final confirmation of the functional destruction of the gene of interest is performed by immunoassays or RNA-based techniques (20). The targeting strategy and the analysis of the integration products by Southern blot hybridization of the human perlecan gene in HCT116 colon carcinoma cells is illustrated in Fig. 2. For the targeting of this gene, a knockout replacenment strategy was considered.

2. Materials

1. Low-electroendoosmosis agarose for electrophoresis (from Fisher).

2. TE buffer: 10 mM Tris-HCl, pH 7.4, 1 mM Na2EDTA.

3. Absolute ethanol (95%), reagent grade.

4. DNA restriction enzymes and related reaction buffers can be purchased from various vendors.

5. Cell electroporation apparatus (example: Hoefer's Progenetor II Electroporation unit).

6. DMEM/FCS: Dulbecco's Minimal Essential Medium with 10% fetal calf serum.

7. DPBS without Ca/Mg: Dulbecco's phosphate buffer saline without calcium and magnesium.

8. Trypsin solution: 0.05% trypsin, 0.53 mMNa4EDTA in Hank's phosphate buffer without Ca/Mg.

Fig. 2. (A) Strategy for targeting of the human perlecan gene in HCT 116 colon carcinoma cells by replacement. The restriction site map of the wild-type allelic locus is illustrated at the top. The solid lines represent the targeted DNA, whereas the dotted line correspond to its flanking regions. An in-scale diagram of the targeting vector constructed by in-frame insertion of the neo selectable marker at the Ncol restriction site of exon II of perlecan is also illustrated. Note the presence of the NcoI site within the neo insert that facilitates the RFLP analysis of the recombination products. The genomic DNA of G418-resistant HCT116 cell clones was digested with NcoI, the products separated by agarose gel electrophoresis, and transferred to a nitrocellulose membrane for hybridization with two 32P-DNA probes identified by the shadowed bars in the figure. Panel (B) illustrates the autoradiogram obtained by using the "external" probe flanking the 3' region of the targeted locus whereas panel (C) illustrates the results with the same blot using a "neo" probe spanning the neo DNA. Lane 1, unsuccessful targeting; lane 2, successful targeting identified by the presence of a NcoI restriction fragment of the expected size (4.6 kb) that hybridizes with both probes; lane 3, evidence for a third integration product giving a larger-than-expected restriction fragment likely generated by the rearrangement of the targeting vector and/or of the targeted locus.

9. Plasmid DNA purification kit from Qiagen or Promega.

10. DH5a-competent Escherichia coli bacteria for transfection (from GIBCO-BRL).

11. Sterile cloning rings (from Scienceware).

12. Geneticin G418 (Neomycin sulfate analog) (Life-Technologies).

13. Hygromycin B (Sigma H 0654).

14. TEA buffer (50x): 2 M Tris base, 1 M glacial acetic acid, 0.05 M EDTA (pH 8.0) to 1 L with water.

15. Ethydium bromide: 1% solution in water.

16. Tris-HCl (pH 7.6) saturated phenol: Mix equal volumes of the phenol and the Tris-buffered solution and let the phases separate at room temperature. Store phenol overlayed with the Tris buffer in a dark bottle.

17. SSC buffer (20x): 3 M NaCl, 0.3 MNa-citrate (pH 7.2).

18. SSPE buffer (20x): 3 M NaCl, 0.25 M NaH2PO4, 0.025 M EDTA (pH 7.4).

19. Denhart's hybridization solution (50x): 10 mg/mL Ficoll 400, 10 mg/mL polyvynil-pyrrolidone, 10 mg/mL BSA fraction V.

20. Thermostatic water baths set at 56 and 65°C. Heating blocks set at 37°C.

21. Thermostatic shakers at 55°C and 65°C.

3. Methods

3.1. Vector Cloning and Linearization

Vector DNA is cloned by standard techniques in quantities to achieve a yield of 200-500 ^g. Following purification by ion-exchange chromatography, the vector is linearized by digestion with a suitable restriction enzyme. The linearized vector is then freed of contaminating proteins and recovered by phenol-chloroform extraction followed by ethanol precipitation. The resulting DNA is solubilized in water and, following linearization is used directly for the transfection into mammalian cells. The linearized DNA can be safely stored at -70°C for prolonged periods of time.

1. Transform recA1 mutated E.Coli bacteria (example DH5a) with the targeting vector and plate on agar using a suitable antibiotic for selection. Pick a formed colony and transfer to 250 mL of LB medium. Grow overnight at 35-37°C or until OD600 reaches ~2.0.

2. Collect the bacteria by centrifugation and extract DNA by the alkaline lysis method followed by DNA purification by ion-exchange chromatography. Several companies (including Qiagen and Promega) sell suitable kits for this purpose.

3. Recover the DNA pellet by ethanol or isopropanol precipitation. Carefully evaporate the excess solvent under a sterile hood but do not allow the pellet to desiccate, as this causes DNA to become insoluble.

4. Dissolve the DNA in 500 ^L of sterile water. Measure the DNA amount by spectrophotometry at 260/280 nm. Assess the quality of the purified DNA by running 1 ^g on 0.7% agarose in TEA buffer. Ethydium bromide-stained DNA should appear as a single band corresponding to supercoiled DNA, with minimal or no nick DNA product evident.

5. Aliquot 100 ^g of DNA into a sterile tube. Bring the final volume to 500 ^L with water and 10x incubation buffer, and incubate for a minimum of 3 h with 50 U of a suitable restriction enzyme at the manufacturer's suggested temperature (see Note 1).

6. Stop the incubation by heat-inactivating the enzyme. Bring the sample to room temperature and add 250 ^L of saturated phenol solution. Mix by tube inversion several times. Avoid vortexing as this may cause DNA shearing. Add 250 ^L of chloroform and mix carefully. Centrifuge at 5000g for 5 min at room temperature and transfer the supernatant to a new tube.

7. Add 500 ^L of chloroform and mix gently. Centrifuge and transfer the supernatant to a new tube.

8. Add one-tenth of the volume of 3 M Na-acetate (pH 7.6) and 1 mL of ice-cold ethanol. Mix by tube inversion and place the tube at -20°C for no less then 1 h.

9. Recover the precipitated DNA by centrifugation at 10,000g for 20 min in a refrigerated centrifuge. Discard the supernatant and carefully remove the residual solvent.

10. Dry the DNA pellet under a sterile hood and solubilize in sterile water by incubating overnight at 4°C.

11. Measure the DNA concentration by spectrophorometry and assess the quality of the linearized DNA by agarose electrophoresis. Store at -20°C for up to a week.

3.2. DNA Transfection by Electroporation

The initial step of the gene targeting process is the introduction of DNA into the recipient cells. This can be achieved through several means, including DNA electroporation, lipofectamine-mediated, and calcium phosphate-based procedures. In our hands, DNA electroporation achieves the highest degree of transfection efficiency, as evidenced by the number of drug-resistant colonies recovered.

1. Prepare the linearized targeting vector by the procedure described above.

2. One day before the electroporation, passage 1:2 the actively growing cell (at this point ~80% conflent).

3. Feed the cells with fresh medium 4 h before harvesting and electroporation.

4. Wash the plates twice with PBS and detach the cells by treatment with trypsin solution for 10 min at 37°C.

5. Stop the action of the trypsin solution by adding 1 volume of fresh medium and dissociate the cell clumps by pipetting.

6. Centrifuge the cell at 1100g(1000 rpm) for 5 min in a clinical centrifuge and discard the supernatant. Resuspend the cell in 10 mL of electroporation buffer and determine the number of cells.

7. Withdraw and recover by centrifugation 107 cells. Discard the supernatant and resuspend the cell in 1 mL of electroporation buffer.

8. Mix 50 ^g of the linearized targeting vector with the cell suspension in an electroporation cuvette. Incubate for 5 min at room temperature (see Note 2).

9. If HCT116 cells are used, electroporate at 230 V, 1080 ^F, 1 sec. Incubate for 5 min at room temperature.

10. Plate the entire content of the cuvette into 10-cm tissue culture plate with DMEM.

11. Apply G418 selection 24 h after the electroporation.

12. Refeed the cell when the medium starts to turn yellow, usually daily for the first 5 d.

13. Ten days after the electroporation, most of the colonies have reached a size suitable for subcloning (see Note 3).

3.3 Recovery and Expansion of Stable Transfected Colonies

Colonies that have reached suitable degree of growth are processed for DNA extraction and subsequent analysis of the restriction digestion fragments by Southern blot hybridization analysis. A main consideration in the design of the screening proce dure is the possible differential growth of the clones. This implies that the colony harvesting be performed at subsequent times (see Note 4).

1. Wash the plate containing the colonies with Ca/Mg-free PBS.

2. By holding the plastic dish against a light source, visually inspect the bottom. Colonies appears as translucent areas of 2-4 mm in diameter. Circle them with a marker for easy identification.

3. Remove the medium and wash the plates once with DPBS without Ca/Mg. Carefully remove the washing buffer.

4. Streak the bottom of a sterile cloning ring into silicone grease and place it around the colony of interest. The purpose is to seal out the colony.

5. Add 20 ^L of trypsin solution to each cloning ring and place in a humidified incubator for 15 min.

6. Add 50 ^L of media to each ring and transfer the cells to a 48-well plate into which 250 ^L of media per well had been added in advance.

7. Carefully aspirate all the remaining cells. Remove the cloning ring and replenish the dishes with fresh medium containing the selection agent. This allows slowly growing colonies to reach suitable size. Perform a second round of harvesting a week later.

8. Grow the cells that had been transferred to 48-well plates in the presence of selection medium.

9. When the cells are approaching confluence, wash with Ca/Mg-free PBS and dissociate with 50 ^L of trypsin. Add 350 ^L of fresh medium and pipet vigorously to dissociate the cells. Transfer 200 ^L to a 6-well plate and add 2 mL of medium. Once grown, these cells may be stocked frozen. The remaining of the cells are transferred to 12-well plates and expanded. Upon reaching confluence, the clones are processed for DNA extraction.

3.4. Genomic DNA Isolation and Southern Blot Analysis

Cell growth is examined under a light microscope and clones are processed when they have reached 75-100% confluence. Following cell lysis, DNA is precipitated with ethanol and digested with the selected restriction enzyme. The generated restriction fragments are isolated by agarose gel electrophoresis and hybridized with a suitable 32P-labeled probe.

1. Grow the cells to confluence. Cells at this stage are also recognizable by the yellow medium appearing 24 h after medium replacement.

2. Wash the cells with PBS lacking Ca/Mg and add 300 ^L of cell lysis buffer.

3. Incubate at 37°C in an incubator for 10 min and then transfer to an Eppendorf tube. Cell lysate can be stored at -70°C without appreciable degradation of DNA for several years (see Note 5).

4. When sufficient numbers of samples have been collected, add 10 ^L of protease (10 mg/mL) and place the sample in 65°C heat-block overnight.

5. Perform a phenol-chloroform extraction with 0.5 mL of phenol followed by a final extraction with chloroform.

6. Precipitate DNA by adding 2 vol of absolute ethanol. Turn the tube upside down several times until a filamentous DNA precipitate appears. Collect the DNA by swirling a glass rod inside the tube and transfer the DNA to a new Eppendorf tube containing 100 ^L of TE (see Note 6).

7. Let DNA dissolve at 4°C overnight and then store at -70°C.

8. Withdraw a 20-^L aliquot of DNA solution and digest with 20 U of restriction enzyme in a final volume of 60 ^L at the proper temperature overnight.

9. Heat the sample at 65°C for 15 min and then add 10 ^L of sample buffer.

10. If the expected size of the informative restriction fragment is between 2 and 7 kb, apply the sample to a 0.75% agarose gel containing ethydium bromide. For resolution of fragments of different size, adjust the agarose content accordingly within the 0.5% to 1.5% limits. Load the sample in 0.8-cm-wide wells and run at 1.5 V/cm of running gel overnight (see Note 7).

11. Stop the electrophoretic run and take a picture of the gel under a UV light source, taking care to place a fluorescent ruler to the side of the gel to identify the relative migration of the DNA standard ladder.

12. Process the agarose gel for Southern blot and 32P-probe hybridization with cross-linking to the nitrocellulose membrane by UV. Prehybridization is performed at 65°C in 2x SSPE, 1% SDS, and 10% Denhart's solution for 4-16 h. Hybridization is carried out by exchanging the prehybridization cocktail with a fresh solution containing 50-100 x 106 cpm 32P-labeled DNA probe and incubating for 16 h at 65oC. Excess probe is washed out by incubation in 2% SSC, 1% SDS solution at 65oC twice for 15 min each, followed by autoradiography.

4. Notes

1. If the process of linearization gives rise to two DNA fragments, the targeting vector can be recovered following separation by agarose gel electrophoresis. For this purpose use low-temperature melting agarose and recover the DNA by agarase treatment or by ion exchange using glass beads available as part of a kit from Bio101 or other suppliers.

2. Carefully collect all the cells that remain attached to the electrodes, after the electric discharge. This is done by pipetting the medium against the electrodes and collecting the cells in a dish. Cell clumps are dissolved by vigorous pipetting and distributed in four 10-cm plastic dishes containing 13 mL of medium. It is recommended that the dishes with the medium be preincubated for at least 1 h before the addition of the cells, in order to allow the deposition of the collagen and other extracellular matrix macromolecules at the bottom of the dish to facilitate cell adherence.

3. Drug-resistant colonies should be collected at a sufficient stage of growth to maximize the chance of attachment and growth in the cloning dishes. A minimum size of 2-4 mm is recommended. With time, colony density become too high, with the risk of contamination during ring cloning. Furthermore, some overgrown colonies may undergo apoptosis.

4. Cell colonies should be expanded gradually by initial grow in 48-well plates. Grow rate can vary considerably, and care should be taken to trypsinize and replate the cells if signs of growth resting become evident. Usually, cells reaching confluence in 48-well plates are passed to 12-well plates and later to 6-well plates in 1-2 wk.

5. Cells extracts are stored at -70°C for up to 1 yr. Protease is added only at the time of incubation and DNA precipitation.

6. If the DNA amount is low, the addition of ethanol will not result in the formation of visible filamentous material. In this case DNA can be recovered by mild centrifugation at room temperature.

7. Digested DNA is separated onto a 20-cm long agarose gel in TBE buffer at 1.5 V/cm. Forced cooling is not required. Completion of the electrophoretic run requires 16-20 h.

References

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Healthy Chemistry For Optimal Health

Healthy Chemistry For Optimal Health

Thousands Have Used Chemicals To Improve Their Medical Condition. This Book Is one Of The Most Valuable Resources In The World When It Comes To Chemicals. Not All Chemicals Are Harmful For Your Body – Find Out Those That Helps To Maintain Your Health.

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