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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Curr Protoc Pharmacol. Author manuscript; available in PMC 2010 July 14.
Published in final edited form as:
Curr Protoc Pharmacol. 2009 June 1; 45: 14.12.1–14.12.26.
PMCID: PMC2904083

Xenograft and Transgenic Mouse Models of Epithelial Ovarian Cancer and Non Invasive Imaging Modalities to Monitor Ovarian Tumor Growth In situ -Applications in Evaluating Novel Therapeutic Agents


Epithelial ovarian cancer (EOC) is the most commonly fatal gynecologic malignancy in developed countries. Most EOC patients are diagnosed at advanced stage when disease has spread beyond the ovary. While many patients initially respond to surgery and chemotherapy, the long term prognosis is generally unfavorable, with recurrence and development of drug resistant disease. There is a critical need to identify new therapeutic agents that prolong disease-free intervals and effectively manage recurrent disease. Murine models of ovarian carcinoma are excellent models to study tumor biology in the search for new treatments for EOC. Described in this unit are methods for establishing xenograft or allograft models of EOC using ovarian carcinoma cell lines, in vivo imaging strategies for detection and quantification of EOC in transgenic and in xenograft/allograft models, and procedures for necropsy and pathological evaluation of experimental animals.

Keywords: Epithelial ovarian cancer (EOC), xenograft, orthotopic, transgenic mice, metastasis, in vivo imaging, Magnetic Resonance Imaging, Bioluminescent Imaging


The occurrence of spontaneous ovarian carcinoma in experimental animals is relatively rare with a notable exception being the aging hen, where the frequency is ~24% (Fredrickson, 1987). Differences in the reproductive tract anatomy and disease development between chickens and humans limit the use of the aging hen as an animal model for EOC. Many in vivo studies utilized s.c./i.p. (Freedman et al., 1978; Hamilton et al., 1984; Hamilton et al., 1983; Massazza et al., 1989; Ward et al., 1987) as well as orthotopically implanted (Bao et al., 2002; Fu & Hoffman, 1993; Kiguchi et al., 1998) xenografts of human ovarian carcinoma cell lines. Although s.c. tumor formation can be readily quantified with calipers, i.p. or orthotopic implantation of ovarian carcinoma cells provides a more relevant tumor microenvironment. Thus, tumor ‘take’ is less efficient with s.c. injected mice (Kiguchi et al., 1998). Models of ex vivo transformation of rodent, rabbit and human ovarian surface epithelium (OSE) have been described (Adams & Auersperg, 1981; Auersperg et al., 1995; Coppola et al., 1999; Godwin et al., 1992; Liu et al., 2004; Resnicoff et al., 1993; Roby et al., 2000; Testa et al., 1994). * Spontaneously transformed rodent, rabbit, or human OSE-derived cell lines can be grown as xenografts in immunocompromised Nod/SCID or athymic nude recipient mice (Adams & Auersperg, 1981; Coppola et al., 1999; Davies et al., 1998; Godwin et al., 1992; Liu et al., 2004; Roby et al., 2000; Testa et al., 1994) or as tumor allografts in syngeneic rats (Rose et al., 1996) or mice (Roby et al., 2000).

Overview of TgMISIIR-TAg GEM Model of Ovarian Cancer and the Utility of Non-invasive In vivo Imaging Technologies

Despite available technology for making genetically engineered mouse (GEM) models, their use in studying of EOC has been limited by the lack of understanding of the epithelial precursor cell, as well as the critical molecular alterations that contribute to disease initiation and progression (see Background Information). A transgenic model of spontaneous EOC was created by expressing the early region of the Simian Virus 40 genome, including the small and large T antigen (TAg) genes, under transcriptional control of the Müllerian inhibiting substance type II receptor gene promoter (Connolly, 2003; Hensley et al., 2007). Female TgMISIIR-TAg-DR26 transgenic mice develop bilateral ovarian tumors with 100% penetrance by five to six months of age (Hensley et al., 2007) and The tumor histology resembles serous carcinomas, the most frequently diagnosed histologic subtype of clinical EOC. While TgMISIIR-TAg-DR26 mice (Hensley et al., 2007) and other conditional GEM models (Clark-Knowles et al., 2007; Dinulescu et al., 2005; Flesken-Nikitin et al., 2003; Wu et al., 2007) hold promise as preclinical models of human EOC, the time it takes for ovarian tumors to develop is variable and the use of tumor formation as an endpoint precludes the analysis of early lesions. In the absence of reliable methods for disease detection and evaluation of therapeutic response, experiments are limited to long-term studies involving large animal cohorts with a predetermined, and somewhat arbitrary, time point for euthanasia and necropsy. Technologies for in vivo longitudinal imaging of GEM models of ovarian cancer significantly facilitate therapeutic studies allowing for quantitative data acquisition, assessment of response (e.g., stable disease or regression) and minimization of the number of experimental animals required for each study.

Tumor formation resulting from peritoneal or orthotopic implantation of ovarian carcinoma cells, or arising spontaneously in the ovary in GEM models, occurs relatively deeply within the peritoneal cavity making quantification of tumor burden difficult if not impossible. In vivo imaging technologies such as Magnetic Resonance Imaging (MRI), ultrasound, micro-positron emission tomography (micro-PET) and computed tomography (CT) may be used to study mice with spontaneous autochthonous ovarian tumor development (e.g., transgenic, conditional or inducible mouse models) in the absence of fluorescent or luminescent reporters. In particular, MRI is an excellent modality for non-invasive tumor detection, acquisition of high resolution images of anatomical structures and for accurate, serial measurement of ovarian tumors in mice over time (Hensley et al., 2007; Mabuchi et al., 2007). Bioluminescent imaging (BLI) or fluorescent imaging (FLI) may be employed for longitudinal in vivo imaging of tumor burden in GEM mice in which tumor formation is accompanied by expression of a light-emitting reporter such as firefly luciferase or green fluorescent protein (GFP). To use BLI or FLI for human or murine ovarian carcinoma cell lines to be used as tumor xenografts or allografts, cell lines can be engineered to express luciferase or GFP by standard methods of transfection or retroviral transduction. In general, BLI sensitivity is superior to FLI and BLI does not require the subtraction of the autofluorescence background signal from the animal (Choy et al., 2003).

Detailed in this unit are strategies for the uses of ovarian carcinoma cells engrafted in recipient mice (Basic Protocol 1) and a transgenic model of spontaneous ovarian cancer. Both of these models can be used for pre-clinical evaluation of therapeutic agents for treating this condition. Methods for longitudinal in vivo MRI and BLI are also described in detail to facilitate pre-clinical studies and the acquisition of quantitative data on tumor burden over time. Described in Basic Protocol 2 is the use of MRI to detect and quantify the size of spontaneous ovarian tumors arising in a GEM model of EOC (e.g., TgMISIIR-TAg-DR26 mice). The use of BLI to monitor tumor growth and progression in mice allografted with ovarian cancer cells transfected with firefly luciferase is detailed in Basic Protocol 3.

NOTE: Prior to commencing with any experiments involving the use of vertebrate animals, investigators are required to be trained in the proper use and care of experimental animals and to obtain Institutional Animal Care and Use Committee (IACUC) approval for a detailed protocol describing all experiments. All IACUC protocols must conform to governmental regulations regarding the humane use and care of laboratory animals.


Xenograft or allograft models have several advantages over other techniques. For example, ovarian carcinoma cells implanted s.c. allow for rapid, relatively high-throughput quantitative tumor formation studies to test the response to therapeutic agents. In this case tumor volume is measured with a caliper. To study tumor growth in a more relevant microenvironment, ovarian carcinoma cells may also be implanted i.p. or orthotopically by intrabursal injection. Like human EOC, most ovarian cancer cells grown following i.p. or orthotopic administration result in widespread peritoneal disease with tumor implants on serosal surfaces and the production of ascites., Metastases to distant sites, such as the lung or brain are uncommon. Although i.p. or orthotopic implantation of ovarian carcinoma cells may provide a more relevant tumor microenvironment, these approaches make quantitative assessments difficult. However, the ability to manipulate cells in vitro to introduce luminescent and/or fluorescent reporter genes overcomes this limitation by making it possible to use BLI or FLI (BLI is discussed in detail in Basic Protocol 3) to obtain quantitative data. Another advantage of xenograft/allograft EOC models is that target genes can be manipulated directlyin tumor cells in vitro, by constitutive or inducible gene expression or RNA interference, to study the in vivo effects of altered gene expression on EOC etiology or progression. Also, xenograft/allograft EOC models in which equal numbers of cells are implanted and with generally predictable disease latency allows for direct comparisons of the effects of gene expression alterations and/or therapeutic treatments in mice. In the protocol below, step by step methods are described for the preparation of ovarian carcinoma cells and for subcutaneous, intraperitoneal and orthotopic (intrabursal) injections..


Ovarian cancer cell line - There are a large number of human and murine ovarian carcinoma cell lines available from commercial and academic sources, with human cell lines available through ATCC, and human and murine cell lines from individual investigators under a Material Transfer Agreement. The choice of cell line most appropriate for a particular study depends on numerous factors including, but not limited to, the histology of the primary tumor from which the cells were isolated, the ability of cells to produce tumors in immunocompromised mice and relative drug sensitivity and/or resistance. There is an extensive literature regarding the use of human ovarian carcinoma cell lines that can be used to guide the design of individual experiments (Connolly, 2003; Dinulescu et al., 2005; DiSaia et al., 1975; Freedman et al., 1978; Hamilton et al., 1984; Hamilton et al., 1983; Ioachim et al., 1975; Kolfschoten et al., 2000; Mabuchi et al., 2007; Molpus et al., 1996; Ozols et al., 1987; Pieretti-Vanmarcke et al., 2006a; Pieretti-Vanmarcke et al., 2006b; Roby et al., 2000; Szotek et al., 2006; Ward et al., 1987; Woods et al., 1979; Xing & Orsulic, 2005a; Xing & Orsulic, 2005b; Xing & Orsulic, 2006; Zhang et al., 2002))

Cell Culture reagents - unless otherwise specified, standard cell culture reagents can be purchased from commercial sources such as Gibco/Invitrogen and Cellgro.

Cell culture medium – examples include DMEM and RPMI

Fetal calf serum (FCS)

Penicillin/streptomycin (P/S)


100× Insulin/Transferrin/Selenium (Invitrogen or Cellgro)

Phosphate buffered saline (PBS), sterile

Ca++ and Mg++ free PBS (optional)

0.25% Trypsin/EDTA

15 ml conical tubes (Fisher Scientific)

0.4% Trypan Blue

Hemacytometer (Fisher Scientific)

Inverted microscope

37°C water bath (Fisher Scientific)

37°C/5% CO2 incubator (Fisher Scientific)

Tissue culture flasks (75 or 175 cm2) (Fisher Scientific)

Immunocompromised (e.g., Nod/SCID) or syngeneic (e.g., C57Bl/6) recipient female mice, 8–10 weeks of age - available from standard commercial sources, including Taconic, Charles River, and Jackson Labs

Sterile 1 cc tuberculin syringes (Fisher Scientific)

26 G needles (Fisher Scientific)

29 ½ or 30 G needles (Fisher Scientific)

Electric clippers (Fisher Scientific or Roboz Surgical Instrument Co., Inc., Gaithersburg, MD)

Alcohol prep pads (Fisher Scientific)

Betadine (Fisher Scientific)

Sterile surgical instruments- includes scissors, forceps, tissue clips, and surgical wound staples (Roboz Surgical Instrument Co., Inc., Gaithersburg, MD)

6-0 Vicryl sutures (Roboz Surgical Instrument Co., Inc., Gaithersburg, MD)

Calipers for tumor measurement (Fisher Scientific)

Surgical tissue adhesive (Nexaband®, Abbot Laboratories, Chicago, IL)

Dissection microscope

Glass bead sterilizer (optional, Fisher Scientific)

Sterile physiologic saline

10 mg/mI ketamine hydrochloride and 1 mg/mI xylazine hydrochloride in sterile saline (Fisher Scientific)

Buprenorphine (Fisher Scientific)

Heating pad or infrared heat lamp (Fisher Scientific)

Liquid Infant Heel Warmer (optional, Fisher Scientific Cat. No. 22-024-646)

Scale (Fisher Scientific)

Preparation of cells for injection

  1. In a 5% CO2 incubator, grow ovarian carcinoma cells to 60–80% confluence in 75- or 175-cm2 cell culture flasks in appropriate cell culture medium (e.g., DMEM) supplemented with 4% FCS, P/S and L-glutamine and 1× insulin/selenium/transferrin. The number of flasks of cells needed depends on the number of cells injected/mouse and the total number of mice to be injected.
    The murine ovarian carcinoma (MOVCAR) cell line MOVCAR 5009 was established from the malignant ascites of a TgMISIIR-TAg transgenic mouse with bilateral ovarian cancer. This cell line is used as an example in this protocol and in Basic Protocol 3 because it can be grown as an allograft in immunocompromised (Nod/SCID or Nude) or in syngeneic C57Bl/6 recipient mice. Alternatively, the commercially available SKOV3-luc-D3 Bioware® cell line (Caliperls, Hopkinton, MA URL) may be used in xenograft studies.
  2. Remove cell culture medium and wash the cells once with sterile PBS. Add 1–2 ml trypsin/EDTA to a T75 flask (2–4 ml for a T175 flask) and rotate the flask so that the trypsin covers the entire surface of the flask. Rotate the flask occasionally, allowing the cells to remain in contact with the trypsin until they begin to detach from the surface of the flask. The total time for digestion is cell line specific, varying from one to several minutes. For particularly adherent cells, incubation at 37°C may facilitate release of the cells. As soon as the cells start to roll off the surface, dislodge the remaining cells by gently tapping the flask against the base of the palm. Rescue cells from trypsinization by adding a 10:1 ratio of cell growth medium. Transfer the cell suspension to a 15-ml conical tube and centrifuge at 450–500 × g at 4°C for 5 min..
  3. Remove the supernatant by aspiration, add 10 ml complete growth mediumand resuspend the pellet to a single cell suspension by pipeting up and down. Centrifuge at 450–500 × g at 4°C for 5 min.
  4. Remove the supernatant by aspiration, resuspend the cells in 10 ml complete growth medium and place them on ice. Determine the total cells/milliliter in the suspension by diluting the cells 1:100 (i.e., 10 μl cells + 980 μl complete medium + 10 μl trypan blue) and count using a hemacytometer.
  5. Determine the total number of cells needed by multiplying the number of cells to be injected/mouse by the total number of mice (e.g., 5 × 106 cells/mouse × 10 mice = 5 × 107 cells).
    It is recommended that enough cell suspension be prepared for the total number of mice to be used plus at least two additional mice to ensure that a sufficient amount of cells is available in the event of a bad injection or loss of volume from dripping. For example, if the number of mice to be injected is ten, prepare enough cells to inject 12 animals (e.g., 5 × 106 cells/mouse × 12 mice = 6 × 107 cells).
  6. Transfer the calculated volume of cell suspension needed for the animals in the study to a fresh 15-ml conical tube. For example, if the cell suspension contains 1 × 107 cells, transfer 6 ml to a fresh tube, centrifuge at 450–500 × g at 4°C for 5 min..
  7. Aspirate supernatant and resuspend the cells to the desired volume in complete growth medium. Alternatively, cells can be resuspended in PBS. In this case Ca++ and Mg++ free PBS is recommended to minimize cell clumping. Note: The cells should be used as soon as possible to inject mice, but may be stored on ice for up to an hour.
    The number of cells required for tumor formation is cell line-dependent and can be determined empirically, from the literature, or by communication with the investigator providing the them. The volume of cells needed is injection site-specific as discussed below.

Implantation of cells in recipient mice

A. Subcutaneous (s.c.) injection

  • 8a. Gently restrain mouse manually and shave a small area (1.5 – 2.0 mm2) of flank with portable electric hair clippers. Return the mouse to a holding cage and shave remaining animals..
  • 9a. When preparing cells for s.c. injection (steps 1–7 above), resuspend the pelleted cells so the desired number of cells will be delivered in a final volume of 0.1 to 0.2 ml.
    If the total number of cells to be injected is 5 × 106 cells, resuspend the cells to 5 × 107 or 2.5 × 107 cells/ml to inject 0.1 or 0.2 ml/mouse flank, respectively.
  • 10a. Gently swirl cells to ensure the suspension is uniform and load the syringe with no needle attached by drawing up the required volume. Attach a sterile capped 26-G needle to the syringe barrel.
    Inddividual syringes should be used for each mouse to ensure that there are no differences in the total cell number injected attributable to settling or “stacking” of cells adjacent to the syringe plunger.
  • 11a. Holding the syringe upright, remove the cap from the 26G needle, tap syringe to dislodge air bubbles and eject any air bubbles by gently depressing the plunger.
  • 12a Pick up shaved mouse and allow it to grab the top of the cage with its front paws. Gently restrain by holding across scapula or by holding the tail and allowing the mouse to stretch.
  • 13a. Swab the shaved region with an alcohol pad to disinfect the skin.
  • 14a. Gently “pinch” the shaved skin between thumb and finger and with the bevel of the needle facing upward and slightly parallel to the skin surface, insert the tip of the needle through the skin to just below the surface and inject 0.1 – 0.2 ml cells subcutaneously into the shaved flank. Be sure to allow enough of the needle to penetrate to ensure it is completely through the skin. If the needle is inserted correctly a bubble should be visible beneath the surface of the skin following injection.
    If no bubble is visible, it is likely that the injection was intra-muscular. For individuals that have not performed this type of injection, it is recommended to initially practice the s.c. injection method using PBS.
  • 15a. Repeat steps 5–7 for the remaining mice.
  • 16a. Weigh mice and monitor the injection site 1–2 times each week for tumor formation. As tumors become palpable, measure the length and width using calipers. Record all weight and tumor measurements. Tumor volume calculations may be made using the following formula: length × width2 × 0.5 (volume measurements are expressed as mm3 or cm3)
  • 17a. Euthanize by CO2 asphyxiation followed by cervical dislocation those animals bearing tumors that reach 10% of the total body weight, that gain or lose >10% of total body weight, or display signs of compromised health, such as lethargy, hunching, ataxia, ruffled fur, anorexia, or vocalizations.

B. Intraperitoneal (i.p.) injection

  • 8b. Prepare cells as described for s.c. injection in steps 1–7 and 9a above.
  • 9b. With the thumb and forefinger pick the mouse up by the loose scruff of the neck and anchor the tail against the base of the palm with the pinky finger. Turn the mouse over to expose the belly and invert the mouse so that it is upside down. Very gently shake downward so that gravity allows the peritoneal organs to rest on the diaphragm.
  • 10b. Swab the region of the abdomen adjacent to the hind leg with an alcohol pad to disinfect the skin. (Note: shaving is optional)
  • 11b. With the bevel of the needle facing upward and the needle at a slight angle to the abdomen, gently insert the needle through the skin between the rear leg and the midline to target the peritoneal cavity and avoid internal organs such as bladder or liver. Turn the needle so the bevel faces into the peritoneal cavity and inject 0.1 to 0.2 ml of cells.
    If a bubble is appear under the skin at the injection site the cells were injected s.c. rather than i.p. and the animal should be removed from the study. For those who have not performed an i.p. injection it is recommended they initially practice it using PBS.
  • 12b. Repeat steps 9b, 10b, and 11b for remaining mice.
  • 13b. Weigh mice and physically exam them 1–2 times weekly. Mice should be euthanized by CO2 asphyxiation if they exhibit any of the signs described in step 17a above, or have obvious abdominal distension indicating the presence of ascites or major changes in pallor around the eyes and feet, suggesting anemia.
    If cells are modified to express a reporter gene, tumor growth can be also be monitored using a non-invasive in vivo method as described in Basic Protocol 2. Imaging should not take the place of physical observation and monitoring of the mice.

C. Orthotopic (intrabursal) injection

  • 8c. One day prior to surgery, visually inspect all mice to confirm their general health.
  • 9c. Prepare cells for intrabursal injection as described in steps 1–7 above except that they should be resuspended so the desired number of cells are delivered in a final volume of 10 – 20 μl.
    Without removal of the ovary, the maximum volume of fluid that can be injected into the intrabursal space is approximately 10 – 20 μl; therefore the maximum number of cells that can be injected in this manner is limited. Because the volume is so small it is technically challenging to measure precisely the volume injected. However, resuspension of the cells to a concentration of 4 × 107cells/ml will allow injection of 4 × 105 – 8 × 105 cells within the intrabursal space.
  • 10c. Gently swirl cells to ensure suspension is uniform and load the syringe with no needle attached by drawing up the required volume. Attach a sterile capped 29½ or 30-G needle to the syringe barrel.
    If the same syringe is to be used to inject two or more mice, pre-load the it with sufficient volume for the total number of mice (n) plus one or two additional animals.
  • 11c. Holding the syringe upright, remove the cap from the needle, tap the syringe to dislodge air bubbles and eject any bubbles by gently depressing the plunger. Set the syringe aside while performing surgery to expose the reproductive tract.
    Conduct all surgical procedures under aseptic conditions. This includes wearing surgical attire, including surgical scrubs and sterile surgical gloves, sterilization of all surgical instruments and removal of hair from, and disinfection of, the operative site.
  • 12c. Anesthetize mice by i.p. injection of 95 μl per 10 gram body weight of 10 mg/mI ketamine hydrochloride and 1 mg/mI xylazine hydrochloride in sterile saline. Verify anesthesia by checking pedal and corneal reflexes. To monitor pedal and corneal reflexes, use forceps to pinch the paw and a moistened cotton swab or gauze pad to gently brush the eye area. If the mouse retracts the paw or blinks, an adequate state of anesthesia has not been attained. Allow more time for the anesthesia to take effect and check reflexes again. Do not proceed until reflexive responses cease.
    Caution for use Ketamine and Xylazine combination: If ketamine and xylazine are used for surgery lasting longer than 20 min., animals may require additional anesthetic. A second dose of ketamine alone may be safer than the drug combination as the cardiovascular depression caused by xylazine, an α2-adrenoceptor agonist, often outlasts the sedation or analgesia produced by the initial administration of this agent (see Alternatively, mice can be anesthetized by inhalation of 1–3% isofluorane in oxygen (up to 5% for initial induction).
  • 13c. Place anesthetized mouse face down (on its abdomen) on a sterile gauze pad with the head facing away from, and the tail facing toward, the investigator.
  • 14c. Depending on which ovary will be injected, shave a small portion of the back to the right or left of the midline (Figure 14.1A) and disinfect the area with a Betadine-soaked sterile gauze pad and then with a sterile surgical alcohol pad.
    Figure 14.1
    (A) Incision site (dotted line) for orthotopic intrabursal injection procedure. (B) Diagram of ovarian fat pad, ovary, fallopian tube and uterus after withdrawal from peritoneal cavity. Arrows indicate relevant structures, including the infundibulum which ...
  • 15c. Using sterile forceps and surgical scissors gently lift the skin and make a small vertical incision in the dorsomedial position directly above the ovarian fat pad (Figure 14.1A). The peritoneal wall should be visible beneath the incision and the ovarian fat pad (round white area contrasting with dark pink tissue surrounding it) should be visible just beneath the surface of the peritoneal wall.
  • 16c. Gently lift the peritoneal wall lining and make a small vertical incision similar to that done for the skin.
  • 17c. Place a sterile saline-soaked gauze pad on the midline adjacent to the incision. While holding the incision open with forceps, use another sterile forceps to carefully pull the ovarian fat pad through the incision toward the midline (Figure 14.1B). Turn the fat pad over slightly to expose the uterus, oviduct and ovary and allow the entire structure to rest on the sterile gauze.
    Extrication of the fat pad in this manner minimizes nerve and blood vessel damage and exposes the ovary and associated oviduct. Attachment of a sterile surgical clip to the opposite side of the fat pad serves as an effective counterweight to hold the tissue in place over the midline.
  • 18c. Using a dissection microscope, identify the oviduct tubule bend that is closest to the bursa and insert the needle of the syringe into this structure (Figure 14.1B). If the needle is inserted close enough to the infundibulum (opening of the oviduct), the needle will be visible underneath the bursa. Slowly depress the plunger to inject the cells into the space between the bursa and the ovary. The total volume of liquid to be injected intrabursally is 10–20 μl. Following injection,, slowly remove the needle and observe the injection site carefully to ensure that the cell suspension does not leak from the bursa. If injected correctly, the bursa should appear slightly plump or distended.
    It is essential to perform this procedure gently and with precision to ensure the oviduct and bursa are not damaged by the needle or that the bursa does not rupture due to excessive pressure or overfilling. In the event the bursa tears, surgical tissue adhesive can be used to seal the tear and prevent leaking of the injected cells.
  • 19c. Gently replace the reproductive tract and fat pad under the body wall in the same position it was found. Close the incision in the peritoneal wall with 2–4 sutures using 6-0 silk or Vicryl sutures. Close the skin with surgical staples or wound clips. Clean the outside of the skin with sterile saline.
  • 20c. To avoid hypothermia and speed recovery, place the mouse back in a cage on an infant heel warmer or under an infrared heat lamp until it has fully recovered from anesthesia.
  • 21c. To minimize pain, treat mice undergoing surgery with buprenorphine (0.05–0.1 mg/kg s.c. every 8–12 hours for 48 hours after surgery).
  • 22c. Monitor mice every 30 min for 2 h after surgery for choking, bleeding, rapid breathing, vocalizations, head tilting or other persistent abnormal behaviors.
    Euthanize mice as necessary (Step 17a above) by CO2 asphyxiation and cervical dislocation.
  • 23c. Monitor the appearance of the incision for signs of infection such as redness, swelling, or discharge, as well as the behavior and mobility of mice once daily for 10 days postoperatively. If infection occurs seek veterinary care immediately. Surgical skin staples can be removed 7 days after surgery.
  • 24c. Monitor mice daily (physically as in step 13b above) for general wellness and symptoms that necessitate euthanasia (step 17a) Also, as in step 13b above, if cells express appropriate optical reporters, non-invasive imaging can also be used to monitor orthotopic tumor growth.


MRI is particularly well suited for imaging GEM models. Both normal mouse ovaries and primary tumors arising in the ovary are generally well circumscribed and can be identified easily using MRI scanners with a field strength of 7 Tesla or greater (Hensley et al., 2007; Mabuchi et al., 2007). Tumor formation arising from ovarian carcinoma cell lines implanted orthotopically by intrabursal injection may also be visualized and monitored using this instrumentation.. MRI is well suited for monitoring spontaneous ovarian tumor growth in vivo because of its superb soft tissue contrast and high spatial resolution

MRI is not as well suited to imaging diffuse peritoneal disease associated with ovarian metastasis or with tumor formation resulting from i.p. injection. For this purpose BLI (as described in Basic Protocol 3 below) should be used.


GEM models of EOC or mice with orthotopically implanted ovarian carcinoma cell lines (see Basic Protocol 1)

1× sterile filtered PBS

Gadolinium-diethylenepentaacetic acid (Gd-DTPA, Magnevist, Berlex Labs, Hamilton, NJ URL)

Access to a vertical or horizontal bore Magnetic Resonance Imaging scanner, field strength 7 Tesla or greater, with 25–30mm birdcage coil for imaging

Plastic tubing (2–3 mm inner diameter) for phantom to mark orientation of mouse

Isoflurane - oxygen based anesthesia system

Induction chamber for anesthetizing mice (Summit Medical Equipment, Molecular Imaging products,

Electric clippers or lotion depilatory

1ml tuberculin syringe

30 G needle

Infrared heat lamp

Temperature regulation equipment for animal (if available)

MRI - image acquisition

  1. Prepare phantom orientation marker: Dilute Gd-DTPA 1:100 with PBS, fill a 2–3 cm length of soft plastic tubing with the solution and then cap the ends of the tubing.
    Alternatively, vitamin E capsules or any marker that produces a strong MR signal, may be used.
    The purpose of the Gd-DTPA marker is to unambiguously define the left and right side of the animal in the MRI image. This is of particular importance in a vertical bore system, where ‘left’ and ‘right’ as defined by the imaging software is determined by the choice of subject position as ‘prone’ or ‘supine’ which can easily be confused. The left and right sides of the mouse can be reversed in several other ways including transformation of datasets to different formats, different display conventions for different image analysis programs, and operator error in setting subject position parameters. For this reason it is critical to establish and adhere to a convention for phantom placement and to refer to the phantom for unambiguous determination of the left and right sides. In mice the left kidney will generally be located in a position posterior to the right kidney, although in animals bearing large tumors the kidneys may be pushed out of position. For simplicity, our laboratory has established the convention of placing the phantom on the left side of the mouse (Phantom = Left side = Low kidney).
  2. Prepare gadolinium for injection: Dilute Gd-DTPA 1:10 in PBS. Injection volume is 0.2 ml. This corresponds to a dose of 0.2 mg/kg Gd-DTPA for a 25-g mouse.
  3. Remove the mouse from its cage and place it in the anesthesia induction chamber. Set isoflurane at 2% until mice are completely anesthetized. Reduce anesthesia to 1%. Keep mice moderately warm with an infrared heat lamp placed 2–3 feet from the induction chamber.
  4. Remove mouse from the induction chamber and inject (i.m.) the animal with 0.1 ml of Gd-DTPA/PBS solution near each scapula (2 separate injections).
    An i.m. injection of contrast agent will have slower decay than an i.p. injection. Also, an i.p. injection may create a large pool of the contrast agent in the imaging field of view, confusing interpretation of the images.
  5. Place the mouse in the imaging coil with the phantom orientation marker on one side of the animal and set anesthesia at 1% isoflurane. Any sensors for physiological monitoring should be placed on the animal at this point. If respiration is monitored, adjust isoflurane levels such that the mouse takes 1–2 seconds between breaths.
  6. Insert the imaging coil into the magnet and perform standard pre-scanning operations. These include tuning and matching the radiofrequency coil to proper resonance frequency and impedance, optimizing shim settings, setting the MRI center frequency, and calibrating radiofrequency transmit strength.
    On the Bruker DRX system, the Paravision software performs most of these tasks in an automated fashion. The exceptions are the tuning and matching of the radiofrequency coil, which must be done manually.
    For a detailed explanation of high spatial resolution MRI, see “Principles of Nuclear Magnetic Resonance Microscopy” (Callaghan 1991). For a review of recent applications of MRI to mouse models, see NMR in Biomedicine-Special Issue: MR of Transgenic Mice, (Wadghiri and Helpern, 2007).
  7. Perform localizer scans in the axial and sagittal planes (Figure 14.2B and C). Typically, the first localizer scan consists of a 2D spin echo pulse sequence in the axial orientation, 20 slices, 1.5mm slice width, 256×128 matrix, minimum TR=600 msec, 1 average, for a total imaging time of 1 min.. After completion of the first localizer, a second localizer is proscribed in a sagittal orientation.
    Figure 14.2
    MRI of a tumor-bearing TgMISIIR - Tag - DR26 mouse. A) Oblique coronal image showing bilateral ovarian tumors (arrows). The phantom orientation marker is not visible in this slice of the scan, but the left kidney (lk) is oriented posterior to the right ...
  8. Proscribe an oblique slice set, approximately in the coronal orientation (Figure 14.2A). The orientation of these images is such that, if possible, the ovaries, kidneys and uterus are visible on the same slices. To image ovaries in a normal mouse, data sets consist of an interleaved multi slice spin echo pulse sequence. Imaging parameters: slice thickness = 0.5 mm, field of view = 2.56 cm, in-plane resolution of 0.1 mm, with 4 signal averages and 12–15 slices, and repetition time (TR) =500–700 msec (minimized for a given number of slices), for a total scan time of approximately nine min.. As ovarian tumors increase in size in vivo, slice thickness can be increased to 0.75 or 1.0 mm and the number of signal averages decreased to reduce imaging time.
    Imaging in the coronal plane provides investigators with a view that corresponds closely to the view on gross dissection. In addition, respiratory motion artifacts are much less pronounced in images in the coronal plane than those made in the axial or sagittal planes.
  9. Using the oblique coronal images as a reference, proscribe a set of axial images through both ovaries or ovarian tumors, employing MR imaging parameters that are identical to those for the coronal scans. These are for a redundant measurement of tumor volume.
  10. Remove the mouse from the imaging system when the scan is completed and warm it gently with a heat lamp before returning it to its cage. The mouse should recover from the anesthesia in approximately 10 min.
  11. Scan mice every two weeks to obtain longitudinal in vivo tumor growth data.
    In some cases, tumor growth is very rapid necessitating weekly imaging.
    Imaging analysis may be performed off line, and not necessarily immediately following image acquisition.

MRI - Image analysis and volumetrics

A large variety of methods and software are available for image and volumetric analyses including software packages freely available on the internet. This allows for off-line analysis of images, and makes redundant measurements convenient so that measurements from different observers can be compared. The following instructions are specific to the analysis of image sets obtained on a Bruker MR imaging console, which are stored in the Paravision format. Images are converted to “analyze” format and volumetric analyses performed using the free shareware programs Bru2Analyzer and MRIcro (Rorden & Brett, 2000). We apply the standard planimetric technique of manually outlining regions of interest (ROIs) in contiguous slices and the volume is computed by adding the total number of pixels in the ROIs together and multiplying the sum by the pixel size and the slice thickness (Hensley et al., 2007).


MR datasets acquired from scanner

Shareware programs: Bru2analyzer and MRIcro (available online, see below)


  1. Download and install shareware programs onto computer workstation:
    1. This program is used to transfer the images from Paravision to “analyze” format
  2. Transfer original datasets from the MR scanner. Datasets can be burned onto a CD or transferred to the computer workstation directly using a Windows FTP program (e.g., misfiles:
  3. Use the Bru2analyzer program to convert Paravision format to “analyze” format (instructions provided with Bru2anz documentation, or on the website)
  4. Read the “analyze” file using MRIcro program (see MRIcro documentation).
  5. Following the MRIcro documentation, display image slices and, using the region of interest (ROI) tool, outline regions of interest corresponding to the ovary or ovarian tumors (see Examples Studies below and MRIcro documentation for details on drawing and saving ROIs).
    It is often convenient to first choose ROI’s in a slice through the center of the tumor (generally the largest region) and continue towards each end. The ROIs for both the left and right ovaries can be assigned on each image slice. However for post analysis, it may be useful to construct independent ROIs for each ovary and save them as separate files.
  6. Save all ROIs for future reference and for comparison of measurements made by different observers.
  7. Calculate the volume (mm3) for each ovary or ovarian tumor by multiplying the voxel volume by the total number of voxels defined in all ROIs for each ovary.
    Formula: (X)(Y)(Z)(nROI)
    X and Y = in-plane voxel dimensions
    Z = section thickness
    nROI = number of voxels defined in all ROIs for each ovary


To obtain quantitative tumor growth data in mice receiving i.p. or orthotopic injections of ovarian carcinoma cells, cells can be transfected or transduced with a firefly luciferase or fluorescent reporter gene and monitored by BLI or FLI. While both methods are suitable for obtaining quantitative data, the use of BLI may be somewhat more advantageous in that it generally provides superior sensitivity relative to FLI without complicating issues of autofluorescence (Choy et al., 2003). Described below are methods for verification of expression of luciferase in ovarian carcinoma cell lines and for BLI of in vivo tumor growth in mice.


Ovarian carcinoma cell lines expressing a luminescent reporter gene, such as firefly luciferase. Cell lines used include the human ovarian carcinoma cell line, SKOV3-luc-D3 Bioware®cell line, available from Caliperls (Hopkinton, MA). Alternatively, any ovarian cancer cell line of choice (as described in Basic Protocol 1) can be modified by standard transfection or viral transduction protocols to express the firefly luciferase reporter gene.

96-well cell culture dish

Ovarian tumor-bearing mice:

  • -Mice xenografted or allografted with ovarian carcinoma cells expressing a luminescent reporter gene (Basic Protocol 1)
  • -Transgenic mice with a spontaneous tumor that expresses a luminescent reporter gene

In vivo bioluminescent imaging system such as the IVIS Spectrum (Caliper LifeSciences)

1× sterile filtered PBS

Luciferin substrate (Caliper LifeSciences)

Isoflurane - oxygen based anesthesia system

Induction chamber for anesthetizing mice (Summit Medical Equipment, Molecular Imaging products,

1ml tuberculin syringes

30 G needle

Infrared heat lamp

Verification of tumor cell luminescence

  1. Prior to i.p. or orthotopic (intraburasal) implantation of tumor cells, in recipient mice, verify expression of the reporter gene (e.g., firefly luciferase) in vitro by serially diluting 2 × 105 cells/ml 1:2 and plating 100 μl in duplicate or triplicate wells of a 96-well plate to final concentrations ranging from 20,000 cells/well to 39 cells/well.
  2. Add 30 μl (30 μg) of a 1 mg/ml stock solution of luciferin dissolved in PBS to each well. Incubate 5 min at 37°C.
  3. Place cells in imaging chamber and acquire images with an exposure time sufficient to detect the luminescent signal while avoiding saturation of the CCD camera. A typical exposure time is 2 min, with a minimum f-stop setting and a 13.1 cm field of view.
  4. Determine the minimum number of cells needed to detect luminescence given an exposure time of 2–5 min and verify that average bioluminescence is proportional to cell number (see Anticipated Results).

BLI – Image acquisition

  • 5. Inject recipient mice i.p. or intrabursally with a luciferase expressing ovarian carcinoma cells as described in Basic Protocol 1.
    The schedule used for imaging is somewhat dependent on the in vivo growth characteristics of the cell line. For rapidly growing cell lines, the authors routinely image mice immediately following the initial injection to verify successful implantation by the presence of bioluminescent signal and weekly thereafter. For cell lines that are likely to require longer growth periods, image acquisition every other week may be sufficient.
  • 6. To prepare animals for imaging, anesthetize mice by placing them in an induction chamber in the presence of a mixture of 1–2% isoflurane and oxygen.
    Alternatively, injectable anesthetics such as ketamine and xylazine (mixture of 100 mg/kg ketamine and 10 mg/kg xylazine) may be used.
  • 7. Prepare mice for luciferase imaging by removing the hair on the ventral and/or dorsal surfaces using portable clippers or with a lotion depilatory.
    Hair removal may be unnecessary when using cell lines with strong luciferase expression and nude mice, or mice with white fur. Imaging of darker pigmented mice such as, C57Bl/6, will likely necessitate hair removal for optimal image acquisition. The optimal orientation for imaging is dependent on the injection site. For mice injected intrabursally, imaging of the dorsal surface is recommended due to the close location of the ovaries to the back of the animal. For imaging i.p.-injected mice, images acquired from the ventral surface may be sufficient. In either case, as disease advances and spreads throughout the peritoneal cavity, acquisition of images from the opposite surface may also be desired and can be accomplished quickly.
  • 8. Begin flow of anesthesia (2% isoflurane and 1 liter/min oxygen) to imaging system.
    The recommended isoflurane concentration for anesthesia during BLI imaging is slightly higher than for MRI. As BLI imaging times are much shorter, this represents no hazard to the animals. The mice must remain sedated for the full duration of imaging to avoid the potential hazard to both the animal and the equipment of escape and loss within the imaging equipment or the camera chamber.
  • 9. Administer the luciferin substrate (2 mg dissolved in 0.2 ml PBS) by i.p. injection.
  • 10. Place mice in the imaging system, orienting them so that light emitted from tumors will pass through as little intervening tissue as possible before detection by the CCD camera.
    Many systems are designed for imaging several mice at once. Care should be taken to identify each mouse with toe clip, ear punch and/or indelible markers to ensure that the identity of each subject in each image is unambiguous. It is crucial for longitudinal imaging studies to be able to precisely identify each animal from one imaging session to the next. Imaging mice in the same position and with the same group of mice from session to session may simplify post-processing.
  • 11. Record the luminescence signal according to instructions from the imaging system manufacturer. First, record a short exposure time (1 sec) and adjust the imaging time so that the signal can be detected easily without saturating the CCD camera (see Anticipated Results). Record a sequence of images to determine the time after injection at which the signal is maximal (usually 10–15 min). After determining the kinetics of the signal emission it is necessary to perform the imaging only at the time of peak signal in subsequent mice.
    For accurate analyses, it is critical to ensure that acquired images are not saturated.
  • 12. Repeat imaging weekly or every other week to allow estimation of tumor burden over time.
    At the time of final image acquisition, and after euthanasia and necropsy, the mice can be imaged ex vivo to detect and localize tumors present on affected peritoneal surfaces and organs and tissues, such as the peritoneal wall, diaphragm, gastrointestinal and reproductive tracts.

BLI – Image analysis

Perform image analysis using the software that accompanies the imaging system (e.g., Living Image Software for IVIS Spectrum).

  • 13. Define the ROI as the region in the bioluminescent image where the signal exceeds a given threshold. The same threshold should be used each time to accurately compare data from different experiments. As an alternative to choosing a particular signal threshold, the signal over the entire abdomen of the mouse can be integrated in animal models of disseminated peritoneal disease (see Anticipated Results).
  • 14. In a system calibrated for absolute quantification of the emitted signal, such as the IVIS Spectrum, the integrated photon flux (photons/sec) will be corrected for different experimental parameters such as exposure time, F-stop, and field of view. If the imaging system is not calibrated absolutely all of these parameters must be kept constant for all imaging sessions, although the exposure time may be adjusted if the BLI signal becomes so intense that the CCD camera becomes saturated. In this case, signal intensities must be corrected for the different imaging times.
    In mice injected i.p or intrabursally, the signal should be confined to the abdomen.
    While total signal intensities (photons/sec) are generally stronger in Nude or white SCID mice, luminescent signal can also be readily detected in pigmented (e.g., C57Bl/6 mice) mice.


To assure that complete assessments are made in the evaluation of potential therapeutics, it is recommended that a complete necropsy report be generated for each animal. In addition to quantitative assessments of gross tumor burden, collection and fixation of tissues and subsequent histopathologic evaluation may provide important information regarding the morphological and molecular characteristics of the tumors, particularly in response to test agents. Formalin fixed paraffin embedded (FFPE) tissue sections may subsequently be stained with various markers of proliferation, apoptosis, angiogenesis and tumor invasion. Additionally FFPE tissue sections may be stained for the expression of relevant signaling proteins that may be affected by chemotherapeutic agents. In some cases, such as RNA-based applications or immunohistochemical detection for some proteins, it may be desirable or necessary to obtain frozen tissue specimens. Therefore, it is recommended that frozen tissues be saved in addition to FFPE tissues.


CO2 for euthanasia

10% Neutral buffered formalin (NBF) (Thermo Scientific, Fisher)Sterile dissection scissors and forceps

Specimen containers

Sterile transfer pipets

15- or 50-mL conical tubes

Necropsy report form


  1. Technical personnel should monitor the health and well being of the mice 2–3 times a week. As animals require euthanasia, necropsy with gross examination should be performed and accompanied by a full written report. IACUC guidelines require that mice are euthanized when they exhibit signs of illness or distress. These include lethargy, fur scruffing, ataxia, pale skin color indicating anemia, anorexia and/or abdominal distension due to ascites, and large tumor size or, in the absence of such symptoms, when total tumor burden attains 10% of the total body weight.
    Example: As transgenic TgMISIIR-TAg-DR26 mice generally weigh 18–20 g, these subjects are euthanized when total tumor volume (including both right and left ovarian tumors) reaches a maximum of 1500 mm3 to ensure that tumor volume does not exceed the 10% body weight cut-off.
  2. Euthanize mice using an IACUC-approved method, such as CO2 inhalation followed by cervical dislocation.
  3. Place the carcass ventral side up, swab abdomen with 70% ethanol and make an incision with surgical scissors from just above the genitalia to the sternum. Tease the skin away from the fascia with forceps or scissor tips and spread the flaps aside exposing the peritoneal cavity.
    In many cases, mice implanted i.p. or intrabursally with ovarian carcinoma cell lines will have bloody ascites present in the peritoneal cavity. Because for many experiments ascites are collected to isolate primary cell lines from malignant cells, it is useful to expose the cavity with the peritoneal wall intact under aseptic conditions.
  4. Optional: If ascites is present, as evidenced by abdominal distention and the visible presence of blood under the peritoneal wall, and requires collection and/or measurement, prepare a sterile collection tube and sterile transfer pipet. Proceed by making a small incision in the peritoneal wall, insert the tip of a transfer pipet and aspirate the fluid and transfer to a sterile 15 or 50 ml conical tube. Continue aspiration until as much fluid as possible is removed and collected.
  5. Lengthen the incision in the peritoneal wall with scissors to expose the cavity and proceed with necropsy. Note the presence, location, size and weight (measure with calipers and weigh) of tumor nodules within the peritoneal cavity. Carefully remove the gastrointestinal tract, noting the presence and extent of tumor nodules on the mesentery. Locate the uterus and begin to tease it away from surrounding tissue with scissors or forceps. Follow the uterine horns toward the kidney to locate the fallopian tube and ovary. Dissect the ovaries away from the kidney and body wall so the entire reproductive tract can be removed. Tease the junction of the uterus and cervix free and snip through the pubic bone on either side of the vagina to remove the intact reproductive tract. If present, measure the size of the ovarian tumors with calipers and note their gross appearance (e.g., color, consistency, presence of hemorrhagic or serous cysts, etc.). Inspect all other organs for the presence and location of tumor nodules/metastases.
  6. Remove all pathologically altered organs, as well as representative specimens of various organs and tissues, including the brain, lung, liver, kidney, spleen, pancreas, intestine and uterus and place in 10% neutral buffered formalin (NBF) or other fixatives as appropriate. Process as appropriate, such as embedding in paraffin, and sectioning and staining with H&E as required for analyses.
  7. Snap-freeze portions of primary tumors and metastases in liquid nitrogen for subsequent preparation of genomic DNA, RNA and protein lysates.
  8. Discard animal carcass.
  9. Prepare a full necropsy report recording strain, identification code, dates of death and birth, tumor source, such as transgenic or allograft/xenograft tumor including cell injection site and date of cell injection, and note all pertinent observations and measurements.


Background Information

In the United States in 2008 it is estimated that approximately 22,000 new cases of EOC will be diagnosed and 15,520 deaths will occur (Jemal et al., 2008). Although the overall incidence of EOC is relatively low, it is the eighth most commonly diagnosed cancer and fifth most common cause of cancer death among American women. Globally, approximately 204,500 new cases are diagnosed and 125,000 deaths occur annually, making it the sixth most common cancer and the seventh most common cause of cancer death among women worldwide (Parkin et al., 2005). These statistics have changed relatively little over the past 30 years and, as there are currently no reliable means of early detection or prevention of EOC, and they are not predicted to improve in the near term.

Although ovarian tumors can arise from stromal tissue and germ cells, those arising in the epithelium account for 85–90% of all ovarian cancers. The initiating cell population for EOC remains somewhat controversial, with evidence suggesting it originates from either the ovarian surface epithelium (OSE), inclusion cysts lined by OSE (Auersperg et al., 1997; Scully, 1995) or alternatively, components of the secondary Müllerian system, including the epithelial cells of the rete ovarii, paraovarian/paratubal cysts, endosalpingiosis, endometriosis or endomucinosis (Crum et al., 2007; Dubeau, 1999). One reason for the lack of certainty regarding tumor origin is that, unlike epithelial cancers arising in other organs, a well-defined disease spectrum consisting of benign, invasive and metastatic lesions has not been identified for EOC. This is largely because the majority of cases are identified at an advanced stage. In addition, common epithelial tumors consist of several distinct histologic subtypes, including serous, endometrioid, mucinous, clear cell, Brenner and mixed histology tumors (Scully, 1995). EOCs are further categorized as benign, borderline (also referred to as low malignant potential tumors) or malignant. The relationship of borderline tumors to invasive cancers is complex. Molecular evidence suggests that for some histologic subtypes, such as mucinous, endometrioid, and clear cell, borderline tumors are likely related to invasive cancers while others, such as serous, are not (Shih & Kurman, 2005).

Early attempts to use transgenic or other genetic engineering approaches to produce murine EOC models largely resulted in strains that develop sex-cord stromal tumors (Dutertre et al., 2001; Kananen et al., 1995; Keri et al., 2000; Kumar et al., 1999; Rahman & Huhtaniemi, 2001; Risma et al., 1995). Later, a number of laboratories made GEM models of EOC by using ex vivo transformation (Orsulic et al., 2002), transgenic (Connolly, 2003; Hensley et al., 2007) and conditional gene expression strategies (Dinulescu et al., 2005; Flesken-Nikitin et al., 2003; Wu et al., 2007). In 2002, Orsulic et al. described ex vivo transformation of p53 null murine OSE in cultured ovarian explants by retroviral transduction with at least two additional oncogenes (e.g., K-Ras, Myc or Akt). Several laboratories used conditional Cre-LoxP-mediated strategies to establish GEM models of EOC by intrabursal administration of Adenovirus-Cre recombinase. Mice with conditional inactivation of both Rb and p53 in the OSE develop serous ovarian carcinomas (Flesken-Nikitin et al., 2003). Mice with conditional inactivation of Pten and conditional activation of mutant K-ras (Dinulescu et al., 2005) or conditional inactivation of both Pten and Apc (Wu et al., 2007) develop EOCs of the endometrioid subtype. As there is currently no existing mouse model that exhibits OSE-restricted expression of Cre-recombinase, tumor induction that relies on Cre-mediated excision of LoxP-flanked sequences requires intrabursal administration of Adenovirus-Cre (Clark-Knowles et al., 2007; Dinulescu et al., 2005; Flesken-Nikitin et al., 2003; Wu et al., 2007).

Critical Parameters

Orthotopic implantation of ovarian carcinoma cell lines by intrabursal injection requires aseptic micro-surgical technique to avoid infection and post-operative complications. Successful implantation of cells is highly dependent on careful and precise handling of the reproductive tract and injection through the infundibulum into the intrabursal space to ensure the bursa does not tear and thereby allow cells to leak into the bursal space. Bursal tears and/or leaking cells from the bursa will effectively result in an i.p. injection.

As for any procedure that involves survival surgery, administration of anesthesia and post-operative monitoring are essential for the well-being of the animals and a successful outcome.

The procedures outlined in this unit involve either spontaneous tumor development or tumor induction by implantation of ovarian carcinoma cell lines. In either case, mice must be monitored regularly to ensure their well being and to assess the need for euthanasia. Mice should be euthanized as soon as they exhibit any of the following symptoms: 1) loss of > 10% of body weight, 2) general signs of ill health (e.g., lethargy, fur scruffing), 3) ataxia, 4) significant ascites accompanied by weight gain of 2 g over several days, 5) anorexia, 6) dehydration, 7) vocalizations indicating pain or distress, 8) severe anemia indicated by pallor of skin on the feet or around the eyes and/or 9) open, bleeding, or infected sores or wounds.


Listed on Table 14.1 are potential problems that may be encountered in generating xenografts or allografts. Tables 14.2 and 14.3 detail potential difficulties with MRI and BLI. Possible causes and recommended actions for overcoming and/or avoiding procedural problems are indicated in each Table.

Table 14.1
Troubleshooting s.c., i.p. or intrabursally injected tumor xenograft or allografts
Table 14.2
Troubleshooting MRI imaging of spontaneous ovarian tumors
Table 14.3
Troubleshooting BLI imaging of tumor xenografts or allografts ovarian tumors

Anticipated Results – Example Studies using Ovarian Carcinoma Models and Non-Invasive Imaging Modalities

Magnetic Resonance Imaging

Representative images of MRI scans in the oblique sagittal orientation of two wild type (top panels) and two TgMISIIR-TAg transgenic (bottom panels) female mice are shown (Figure 14.3). A single section is depicted unmasked (left) and masked (right) for each animal. The masked images highlight the ROI; that is the normal ovary in wild type mice (top right) or ovarian tumors in the TgMISIIR-TAg transgenic mice (bottom panels) for each animal. Note the presence of the phantom marking the left side of the mouse (right side of the image) in the top panels. Our laboratory has used this method to monitor tumor burden over time in previously described chemotherapeutic studies (Hensley et al., 2007; Mabuchi et al., 2007).

Figure 14.3
Representative examples of coronal slices and regions of interest through the ovaries of two wild type (WT, top row) and two transgenic (Tg) TgMISIIR-TAg mice with ovarian tumors (bottom row). Both unmasked and masked (masked images define the ROIs, e.g, ...

Comprehensive analyses of tumor growth rates and responsiveness to a standard cytotoxic chemotherapy regimen, such as cisplatin and paclitaxel, and a molecularly targeted agent, such as the mTOR inhibitor RAD001 (Everolimus) have been performed (Hensley et al., 2007; Mabuchi et al., 2007). While not curative, TgMISIIR-TAg transgenic mice treated with cisplatin and paclitaxel exhibited a significant delay in tumor progression (Hensley et al., 2007). RAD001 treatment resulted in markedly delayed tumor development, ascites production peritoneal dissemination of ovarian tumors in these mice (Hensley et al., 2007; Mabuchi et al., 2007). Collectively, these data allowed the use of statistical modeling to aid in the design of preclinical studies (Hensley et al., 2007; Mabuchi et al., 2007). Simulations from these analyses suggested that therapeutic studies utilizing TgMISIIR-TAg-DR26 transgenic mice in conjunction with MRI can be conducted with as few as 20 animals in each treatment or control group and that groups that differ by +25% in time to acquire a given tumor volume (e.g., 100 mm3), the acquisition time can be distinguished with 80% power and 5% type I error (Hensley et al., 2007).

Bioluminescence Imaging

For a cell line to be useful for in vivo studies, it must produce a sufficiently bright luminescent signal. Prior to initiating in vivo experiments, it is highly recommended that the luminescent signal be verified in vitro. As an example, serially dilute (1:2) MOVCAR 5009 cells transduced with a retroviral luciferase construct (MOVCAR-5009-Luc) and plate them in triplicate wells ranging from 20,000 to 78 cells/well. After plating, 30 μl of a 1 mg/ml stock solution of luciferin diluted in PBS is added to each well and the plate imaged using the IVIS Spectrum (Caliper LifeSciences). The results of this experiment will demonstrate the luminescent signal can be reliably detected with as few as 312 cells (Figure 14.4A), and verifies these cells are suitable for subsequent in vivo experiments. The IVIS Spectrum allows absolute calibration of signal strength (in photons/second) from a given region of interest, and the brightness of a cell line can be calculated by dividing the calibrated signal from a given well by the number of cells in the well (Figure 14.4B). A signal strength of 5 photons/(sec × cell) is sufficiently bright to be used in mouse models. This corresponds to being able to detect 300 cells in a well with a signal-to-noise ratio of approximately 2 for an exposure time of 2 min.

Figure 14.4
Calibration of the luminescence of luciferase-transduced MOVCAR-5009 cells. A) Image of a 96-well plate in which a serial dilution of cells (from left, 20,000, 10,000, 5,000, 2500, 1250, 625, 312, 156, 78, 39 cells/well) has been performed in triplicate. ...

The MOVCAR-5009 cell line expresses and secretes high levels of VEGF protein into the culture medium and, after i.p. injection, rapidly produces bloody malignant ascites in mice (Connolly, unpublished observations). In a pilot study to test the effects of RNA interference of Vegf expression on tumor formation, a previously described (Dickins et al., 2005) retroviral shRNA expression vector MSCV-LTR-miR-30-PIG (MLP) was used to express Vegf-specific shRNA constructs. Several individual Vegf-specific shRNA constructs were PCR cloned and tested. Levels of Vegf RNA and protein expression in MOVCAR-5009 cells co-transduced with pWZL-Luciferase and retroviral shRNA constructs for Vegf or the MLP vector alone were determined by quantitative RT-PCR and ELISA. Using this method, approximately 80% stable inhibition of Vegf mRNA and protein was observed with several Vegf shRNA constructs (data not shown). In a pilot experiment to test the effects of RNAi-mediated knockdown of Vegf in MOVCAR-5009-Luc cells on tumor growth in vivo, MOVCAR-5009-Luc cells co-transduced with Vegf shRNA 593 were compared to the MLP vector. Mice (n=5 mice/group) were injected i.p. with 5 × 106 MOVCAR-5009-Luc + MLP or MOVCAR-5009-Luc + Vegf Sh593 cells (as described in Basic Protocol 1) and subjected to weekly BLI (as described in Basic Protocol 3) to monitor tumor burden over time. Representative images (Day 7 and Day 21) demonstrated a significant increase in signal intensity over two weeks (Figure 14.5A). The ROIs (indicated by the ovals in Figure 14.5A) were defined as the area of the mouse that emitted luminescent signal and was confined to the abdomen of i.p.-injected mice. Total tumor burden was estimated from the integrated photon flux in the ROI (Figure 14.5B). All image analyses were performed using Living Image software provided by the manufacturer. Mice allografted with MOVCAR-5009-Luc + MLP cells were euthanized 21 days post injection while mice allografted with MOVCAR-5009-Luc + Vegf Sh593 cells did not require euthanasia until 29 days post injection. Therefore, expression of the Vegf shRNA in MOVCAR-5009-Luc cells significantly delayed ovarian tumor progression and the accumulation of ascites as compared to the vector control (Figure 14.5C).

Figure 14.5
Bioluminescent imaging (BLI) of ovarian tumors in SCID mice

In conjunction with the Fox Chase Cancer Center Biostatistics Core, it was determined that when using BLI to determine growth rates, sample sizes of 20 animals/test condition (e.g., therapeutic agent, short hairpin RNA or vehicle) should be sufficient to obtain statistically significant results using the following approach. Regression analysis of log transformed tumor area versus time yields a single growth rate for each mouse. The resulting growth rates (n=40) can be compared using the Wilcoxon two-sample test. Since power cannot be determined in advance for this procedure, the Fisher’s exact test should be used. Thus, the 40 growth rates can be sorted and the resulting data submitted to this test, which is predicted to distinguish odds ratios of 5.3 and 1.0, with 80% power and 4.79% type I error.

Time Considerations

The amount of time necessary to perform the implantation of ovarian carcinoma cell lines is dependent both on the injection method chosen and the experience of the investigator. For s.c. injections, shaving the flank and administering the injection takes 2–3 min per animal. For i.p. injections, a reasonably experienced investigator should be able to inject a cage of 5 mice in less than 5–10 min. For either s.c. or i.p injections, a large number of mice (n= 40–60) can be injected in a few hours’ time. Intrabursal injection is more complicated,, requiring survival surgery and great deal more technical skill to execute the injections correctly. A highly experienced individual, such as a person who has made transgenic animals, would likely be able to complete the surgery and injection of one mouse in 20–30 min. It is essential for this procedure that another individual assist with preparation of animals for surgery, administration of anesthesia, application of wound clips and post-surgery monitoring. With assistance, intrabursal injections can be performed on 15–20 mice in one day.

The time required for tumor formation is largely cell line-dependent. For well studied cell lines, information regarding the recommended number of cells for injection and resulting latency of disease formation can be obtained from the literature or individual investigators that routinely utilize the cells. In the absence of such information, it is recommended that these parameters be determined empirically in pilot studies. For example, latency of disease formation after i.p. injection of 5 × 106 MOVCAR cells can vary from 4–16 weeks depending on the individual MOVCAR cell line used.

For studies aimed at assessing the effectiveness of drug candidates or other therapeutic modalities, frequency and routes of administration, and doses should be determined in advance. All experiments should be planned in consultation with a statistician to ensure that the minimum number of experimental and control animals are used while still allowing acquisition of statistically significant data. The use of in vivo imaging for longitudinal quantification of tumor burden is advantageous and should significantly reduce the number of animals needed. The time required to execute a given study largely depends on the frequency, duration and route of administration of the potential drug.

Contributor Information

Denise C. Connolly, Fox Chase Cancer Center, Philadelphia, PA, Phone: 215-728-1004, Fax: 215-728-2741, ude.cccf@yllonnoC.esineD.

Harvey H. Hensley, Fox Chase Cancer Center, Philadelphia, PA, Phone: 215-728-3156, Fax: 215-728-3574, ude.cccf@yelsneH.yevraH.


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