Stable Transfection Of Cultured Cells With Antisense Dna Constructs

Method A. Transfection by Liposomes

Liposome-mediated transfection is relatively safe and has a high efficiency of transfection in cultured cells. It is suitable for both transient and stable transfection of tissues or cells. It mandates smaller amounts of DNA for transfection compared with electroporation and other methodologies. Lipofectin reagents are commercially available (e.g., Gibco BRL Life Technologies). Liposomes are composed of poly-cationic lipids and neutral lipids at an appropriate ratio (e.g., 3:1). The primary principles of this method are: (1) the positively charged and neutral lipids can form liposomes that can complex with negatively charged DNA constructs; (2) the DNA-liposome complexes are applied to cultured cells and are uptaken by endocy-tosis; (3) the endosomes undergo breakage of membranes, releasing the DNA constructs; and (4) the DNA enters the nucleus through the nuclear pores and facilitates homologous recombination or integration into the chromosomes of the cell.

The primary disadvantage of the lipo-transfection method is that if the ratio of lipids-DNA and the cell density are not appropriate, severe toxicity will be generated, killing quite large numbers of cells. Embryonic stem (ES) cells are more sensitive to toxicity compared with other fibroblasts such as NIH 3T3 and mouse STO cells.

1. Harvest cells by gentle trypsinization as described in Chapter 16. Centrifuge the cells at 900 rpm for 5 min and resuspend the cells well in normal culture medium at approximately 106 cells/ml.

Note: Make sure that the cells are suspended gently but well prior to transfection because aggregation of the cells may cause difficulty in the isolation of stably transfected cell clones.

2. In a 60-mm tissue culture dish, seed the suspended cells at about 1 x 106 cells in 5 ml of normal culture medium. Allow the cells to grow under normal conditions for 12 h. Proceed to transfection.

Note: Mouse ES cells grow fast and form multiple layers of clones instead of the monolayers seen in other cell lines. Because of these features, it is not recommended to overculture ES cells prior to transfection because multilayers of ES cells may result in a low efficiency of transfection and a high percentage of potential cell death. Therefore, once a relatively high density of cells has been obtained, transfection can be carried out as soon as the ES cells attach to the bottom of the dish and grow for 6 to 12 h.

3. While the cells are growing, DNA constructs can be linearized to be transferred by the use of a unique restriction enzyme. Protocols for appropriate restriction enzyme digestion are described in Subcloning of Gene and DNA Fragments in Chapter 4.

4. In a sterile laminar flow hood, dissolve 10 to 20 mg DNA in 0.1 ml of serum-free medium (free of antibacterial agents) and dilute the lipofectin reagents two- to fivefold in 0.1 ml of serum-free medium (free of antibacterial agents). Combine and gently mix the DNA-lipids media in a sterile tube. Allow this mixture to incubate at room temperature for 15 to 30 min. A cloudy appearance of the medium indicates the formation of DNA-liposome complexes.

5. Rinse the cells once with 10 ml of serum-free medium. Dilute the DNA-liposome solution to 1 ml in serum-free medium (free of antibacterial agents) and gently overlay the cells with the diluted solution. Incubate the cells for 4 to 24 h under normal culture conditions. During this period, transfection takes place.

6. Replace the medium with 5 ml of fresh normal culture medium with serum every 12 to 24 h. Allow the cells to grow for 2 to 3 days before drug selection.

7. Trypsinize the cells and plate them at a density of about 3000 cells/100-mm dish or T25 flask culture for 24 h at 37°C with 5% CO2. Proceed to drug selection.

Method B. Transfection by Microinjection

Microinjection is one of the highest efficiencies of transfection, although the number of cells that can be injected within a given time is usually limited, depending on personal experience. The equipment for microinjection should include: (1) an injection microscope on a vibration-free table (Diaphot TMD microscope with Nomarski optics is available from Nikon Ltd); (2) an inverted microscope for setting up the injecting chamber and holding the pipette and needle (SMZ-2B binocular microscope is available from Leitz Instruments Co.); (3) two sets of micromanipulators (Leitz Instruments Ltd.); (4) Kopf needle puller model 750 (David Kopf Instruments); (5) a finer needle and a pipette holder; and (6) a Schott-KL-1500 cold light source (Schott).

1. Thoroughly clean a depression slide with teepol-based detergent and extensively rinse it with tap water and distilled water. Rinse it with ethanol and air-dry. Add a drop of culture medium to the depression slide and a drop of liquid paraffin on top of a drop of medium. This is preparation for suspension cells but not fibroblast cells attaching on the bottom of culture dishes or flasks.

2. Assemble all parts necessary for the injection according to the manufacturer's instructions. Carry out pretesting of the procedure prior to injection of the DNA sample.

3. For cultured fibroblast cells attached to the bottom, transfer the cultured cells onto the stage under the microscope. For cell suspensions, carefully transfer the cells to the medium drop using a handling pipette that can take up the cells by suction under the microscope.

4. Prepare microinjection needles with a Kopf needle puller (model 750). Slowly take the DNA sample up into the injection needle by capillary action.

5. Under the microscope, hold the suspended cells with the holding pipette and use the micromanipulator to insert the needle individually into the cell. Slowly inject the DNA sample into the nucleus. The pressure can be maintained on the DNA sample in the needle by a syringe connected to the needle holder. As long as nucleus swelling occurs, immediately remove the needle a little bit away from the cell. The volume injected is approximately 1 to 2 pl. For cultured fibroblast cells attached to the bottom of a culture dish or flask, no holding of the cell with a pipette is needed. Injection can be directly performed into the nucleus of the cell.

6. Transfer the injected cells to one side and repeat injections for other cells. About 40 to 60 cells can be injected in 1 to 2 h, depending on an individual's experience.

7. Carefully transfer the injected cells back to the incubator. Allow the cells to grow for 2 to 3 days prior to harvesting or drug selection.

Method C. Transfection by Electroporation

Detailed protocols are given in Chapter 8.

Method D. Transfection by Retrovirus Vectors

Detailed protocols are described in Chapter 8.


Detailed descriptions are given in Chapter 8.


This characterization will provide information about whether or not the developed clones contain and express antisense-induced inhibition in the cells.

Analysis of Gene Underexpression at the Protein Level by Western Blotting

The detailed protocols for protein extraction and western blot hybridization are described in Chapter 11, Analysis of Gene Expression at the Protein Level. The major focus of this section is on how to analyze western blot data. Assuming that the product or protein of the cDNA is 80 kDa and that monoclonal antibodies have been raised to be against the 80-kDa protein (shown in Figure 9.4), a strong 80-kDa band is detected in control cells (lane 1). In contrast, putative antisense clones show a significant reduction of the 80-kDa protein (lane 2 to lane 4) compared with the control cells.

Examination of Expression of Antisense RNA by Northern Blotting

The detailed protocols for RNA isolation and northern blot hybridization are described in Chapter 11. The major focus of this section is on how to analyze northern blot data. Assuming that the antisense RNA of the gene is 0.8 kb and that a specific sense RNA probe is used (shown in Figure 9.5), no band is detected in control cells (lane 1). However, four stably transfected clones clearly show expression of antisense RNA (lane 2 to lane 5).

116 80

FIGURE 9.4 Protein analysis of antisense clones by western blotting. Lane 1: nontransfected cells. Lane 2 to lane 4: underexpression of the 80-kDa protein in different antisense cell clones.

' Protein of interest

FIGURE 9.4 Protein analysis of antisense clones by western blotting. Lane 1: nontransfected cells. Lane 2 to lane 4: underexpression of the 80-kDa protein in different antisense cell clones.

' Protein of interest

Antisense RNA

FIGURE 9.5 Antisense RNA analysis by northern blotting using a sense RNA probe. Lane 1: nontransfected cells. Lane 2 to lane 5: expression of antisense RNA (0.8 kb) in four antisense clones.

Determination of Integration Copy Number by Southern Blot Analysis

Chapter 7 describes the detailed protocols for Southern blot hybridization. Assuming that genomic DNA is digested with a unique restriction enzyme that cuts outside the endogenous gene or cDNA introduced and that the cDNA is labeled as the probe for hybridization (shown in Figure 9.6), lane 1 is the control cells and contains one band. Lane 2 and lane 4 have one copy of integration of the introduced cDNA. Lane 3 is a clone that shows two copies of insertion of the cDNA.

Expression Assay of Reporter Genes

Detailed protocols are described in Chapter 8.


There are two methods for the production of transgenic mice. One is to develop stably transfected mouse ES cells with antisense cDNA constructs; these cells can be injected into female mice to generate genetically altered animals. The other method is to inject antisense cDNA constructs directly into mouse oocytes by microinjection. The injected oocytes are then transferred into the oviduct of a female mouse to produce transgenic mice.

FIGURE 9.6 Antisense DNA analysis by Southern blotting using the introduced cDNA as a probe. Lane 1: nontransfected cells. Lane 2 and lane 4: single-copy integration of antisense cDNA. Lane 3: two copies of insertion of antisense cDNA.

Method A. Production of Transgenic Mice from Stably Transfected ES Cells

Selection of C57BL/6J Estrous Females

The C57BL/6J mouse is a well-established strain widely chosen as the best choice of host embryo for ES cell chimeras and germline transmission. These mice have a coat color different from 129 SVJ mice that can be used as a coat color selection marker. In order to synchronize the estrous cycle relatively and simplify female selection, it is recommended that female mice be caged together without a male. Their estrous cycle is approximately 3 to 4 days and ovulation usually occurs at about the midpoint of the dark period in a light/dark cycle that can be altered to meet a schedule. For example, the cycle may be adjusted to 14 h of light and 10 h of darkness. In this way, most strains of mice produce blastocysts that can be utilized for injection in the late morning of the third or fourth day.

The question here is how to identify pre-estrous or estrous females for mating. The basic criteria are the external vaginal epithelial features. At the pre-estrous stage, the vaginal epithelium is usually characterized by being moist, folded, pink/red and swollen. At the estrous stage, the vaginal epithelium, in general, is dry but not wet, pink but not red or white, wrinkled on upper and lower vaginal lips, and swollen. After the females have been caged for a while and when they reach the prediction of pre-estrous stage, they should be monitored on a daily basis. Once they are at the estrous stage, mating should be allowed to occur.

An alternative way to promote the estrous cycle is through hormonal regulation of ovulation with exogenous hormones — a procedure termed superovulation. Injection of pregnant mice with mare serum gonadotropin (PMSG, Sigma) can stimulate and induce immature follicles to develop to the mature stage. Superovulation can be achieved by further administration of human chorionic gonadotropin (HCG, Sigma). The procedure is outlined as follows:

1. Three days or 72 to 75 h prior to mating, induce superovulation of 10 to 15 female mice (3 to 4 weeks old) by injecting 5 to 10 units PMSG intraperitoneally into each female between 9:00 a.m. and noon on day 1.

2. Two days or 48 to 50 h later, inject 5 to 10 units of HCG intraperitoneally into each female between 9:00 a.m. and noon on day 3. After injection, cage the females and proceed to mate them with sterile males.

Tip: Superovulation will occur about 12 h following HCG administration. The injection time at step (b) will coincide with the midpoint of the dark cycle.

Preparation of a Bank of Sterile Males by Vasectomy

While females undergo their estrous cycle, prepare a stock of sterile males to mate with the estrous female chosen as the embryo transfer host so as to generate pseudopregnancy. The following procedures are recommended to be performed in conditions as sterile as possible.

1. Weigh a mouse and use 0.2 to 25 ml of tribromoethanol to anesthethize it. Swab the abdomen with 70% ethanol using cotton.

2. Lift the skin free of the peritoneum with forceps and carefully make a 1-cm transverse skin incision at approximately 1 cm rostral to the penis. Carefully make another similar incision through the peritoneum.

3. Use blunt forceps and reach laterally into the peritoneum and carefully grasp the testicular fat pad located ventral to the intestine and attached to the testis. Through the incision, gently pull out the pad together with the testis.

4. Distinguish the vas deferens from the corpus epididymis and carefully separate the vas deferens from the mesentery by inserting the closed tips of the iridectomy scissors through the mesentery. Then, open the scissors and leave them at this position to support the isolation of a length of the vas deferens.

5. Insert two pieces of suture under the vas deferens with forceps, tie up the vas deferens and, with the forceps, at two sites approximately 2 cm apart, cut out a piece of the vas deferens (about 1 cm in length) between the two knots.

6. Return the testis to the peritoneum and perform the same procedures on the other side.

7. Suture the peritoneal wall and clip the skin incision with a couple of small wound clips. Leave the vasectomized male for 10 to 14 days to clear the tract of viable sperm and to ensure that he is sterile prior to use for mating.

Materials Needed

Small blunt, curved forceps Three pairs of watchmaker's forceps Small sharp scissors Iridectomy scissors

Note: These surgical instruments can be wrapped in foil and kept in a 120

to 140oC oven for at least 6 hours prior to use. Surgical gloves Small wound clips Tribromoethanol anesthesia 70% Ethanol and cotton Surgical silk suture and needle

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