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In vivo Efficacy Studies in Cell Line and Patient-derived Xenograft Mouse Models
在细胞系和患者源性异种移植瘤小鼠模型中进行的体内疗效研究   

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Abstract

In vivo xenograft models derived from human cancer cells have been a gold standard for evaluating the genetic drivers of cancer and are valuable preclinical models for evaluating the efficacy of cancer therapeutics. Recently, patient-derived tumorgrafts from multiple tumor types have been developed and shown to more accurately recapitulate the molecular and histological heterogeneity of cancer. Here we detail the procedures for developing patient-derived xenograft models from breast cancer tissue, cell-based xenograft models, serial tumor transplantation, tumor measurement, and drug treatment.

Keywords: Patient-derived xenograft(患者源性异种移植瘤), Tumor transplantation(肿瘤移植), Mammary fat pad(乳房脂肪垫), Tumor measurement(肿瘤测量), Dosing(剂量)

Background

Xenograft models have served as a robust method for investigating the genetic drivers of cancer and determining the potential efficacy of cancer therapeutics. The ability to propagate human cancer cells and tissues in mice was drastically advanced with the discovery of T-cell deficient athymic nude (nu/nu) mice and T- and B-cell deficient severe combined immunodeficient (scid/scid) mice (Flanagan, 1966; Bosma and Carroll, 1991). Since these discoveries additional immunocompromised mouse models have become available including, recombination-activating gene 2 (Rag2)-knockout mice, non-obese (NOD)-scid mice, and NOD-scid IL2Rgamma(null) mice (also known as NSG mice) (Shinkai et al., 1992; Prochazka et al., 1992; Shultz et al., 2005). These immunocompromised mouse models have enabled the development of numerous and diverse in vivo models of human cancer.

There are several options that should be considered when developing a xenograft model including the site of injection or implantation. Subcutaneous xenografts are often used in in vivo studies due to tumor accessibility for growth measurement and imaging; however a significant limitation of this model is the lack of a normal stromal microenvironment for most cancer cells. Orthotopic xenografts offer a complementary stromal microenvironment; however there are also disadvantages to this route depending on the orthotopic site, including more complex surgical procedures, difficulty of measuring tumor growth or response, and the limitations of rodent stroma (Talmadge et al., 2007). The use of orthotopic xenografts has been used extensively in many cancer studies, especially breast cancer research. Injecting into the mammary fat pad is a relatively simple procedure that allows for the visible and measurable growth of breast cancer cells. Even though the mammary fat pad offers a complementary tissue site for breast cancer cells, it is important to note there are distinct differences between the human and rodent mammary stroma and hormonal environment.

Xenograft models have been used extensively as predictive models of cancer therapeutic efficacy. For preclinical studies, it is essential to evaluate drug efficacy and potential toxicities in vivo. Even though in vivo preclinical studies are valuable, the results have not consistently translated to the clinic and the significance of these studies are debated (Talmadge et al., 2007; Sausville and Burger, 2006). There are several variables that need to be considered when designing drug studies such as the appropriate cell lines (or PDX models), dosage and dosing schedules, and statistical analysis. Each of these factors should be carefully considered in order to most closely mimic human cancer progression and treatment response.

Recently there have been significant advances in the development of patient-derived tumor xenografts (PDX). PDX models have the advantage of maintaining the molecular and histological heterogeneity of the original tumor (DeRose et al., 2011). Moreover, they have been shown to be superior at predicting drug response compared to standard cell culture xenograft models (Hait, 2010; Fruchter et al., 1990; Voskoglou-Nomikos et al., 2003; Gao et al., 2015). Recent studies have advanced the success of establishing breast cancer PDX models that recapitulate the molecular, stromal, and phenotypic heterogeneity that exists in breast cancer (DeRose et al., 2013). Overall, cell line-based and patient-derived xenografts are essential models for investigating cancer initiation, progression, and treatment response. Here, we describe the protocols for developing cell-based xenograft models, patient-derived xenograft models from breast cancer tissue, serial tumor transplantation, tumor measurement, and drug treatment.

Materials and Reagents

  1. 25 gauge needle (for inoculations) (BD, catalog number: 305127 )
  2. Tissue culture-treated culture dish (Corning, catalog number: 430293 )
  3. Eppendorf tubes (1.5 ml) (Eppendorf, catalog number: 022363204 )
  4. Falcon 15 ml conical centrifuge tubes (Corning, catalog number: 352196 )
  5. Cryogenic vial
  6. Trocar syringe, 10 gauge (Innovative Research of America, catalog number: MP-182 )
  7. 1 cc U-100 insulin syringe, 28 gauge x ½” needle (for Ketoprofen injections) (BD, catalog number: 329410 )
  8. Oral gavage tips, 22 gauge x 1” with 1 ¼ mm ball (Cadence, catalog number: 7901 )
  9. Tuberculin syringe, 1 ml (for inoculations) (BD, catalog number: 309659 )
  10. Betadine solution swab sticks (Thermo Fisher Scientific, Fisher Scientific, catalog number: 19-061617 )
  11. Sterile alcohol prep pads, 70% isopropyl alcohol (Thermo Fisher Scientific, Fisher Scientific, catalog number: 22-363-750 )
  12. Sterile cell preparations or patient-derived tumor tissue
  13. 4-7 week old Athymic Nude mice [Crl:NU(NCr)-Foxn1nu] (Charles River Laboratories International, catalog number: 490 )
  14. 4-7 week old NSG mice (NOD scid gamma, NOD-scid IL2Rgnull, NOD-scid IL2Rgammanull) (THE JACKSON LABORATORY, catalog number: 005557 )
  15. Trypsin or appropriate enzymes
  16. Hanks’ balanced salt solution modified (HBSS) (Thermo Fisher Scientific, GibcoTM, catalog number: 14170112 )
  17. Research Animal Diagnostic Laboratory (RADIL) infectious microbe PCR amplification test (IMPACT) (IDEXX BioResearch)
  18. Phosphate buffered saline (PBS) pH 7.4 (Thermo Fisher Scientific, GibcoTM, catalog number: 10010049 )
  19. Penicillin-streptomycin (5,000 U/ml) (Thermo Fisher Scientific, GibcoTM, catalog number: 15070063 )
  20. 70% ethanol
  21. Surgical cleaner
  22. Isoflurane liquid inhalant OD 99.9% (Henry Schein Medical, catalog number: 1182097 )
  23. Ketofen® (Ketoprofen) (Patterson Veterinary Supply, catalog number: 10004029 )
  24. Absorbent underpads (Thermo Fisher Scientific, Fisher Scientific, catalog number: S67011 )
  25. 10% neutral formalin (Sigma-Aldrich, catalog number: HT501128-4L )
  26. FBS
  27. DMSO
  28. Optional Materials and Reagents
    1. 17β-Estradiol, 60-day release pellets (Innovative Research of America, catalog number: SE-121 )
    2. Veterinary grade surgical glue (Patterson Veterinary Supply, catalog number: 07-805-5031 )
    3. Cryogenic media (see formulation below)
    4. Cell freezing container (Corning, catalog number: 432010 )
    5. Internal thread cryogenic vials, 2.0 ml (Corning, catalog number: 430488 )
    6. Liquid nitrogen

Equipment

  1. Digital calipers (Thermo Fisher Scientific, Fisher Scientific, catalog number: 06-664-16 or Mitutoyo America, catalog number: 500-163-30 )
  2. Safety scalpel (Merit Medical Systems, catalog number: SMS210 )
  3. Wound clip applier 9 mm (Roboz Surgical Instrument, catalog number: RS-9260 )
  4. 9 mm wound clips (Roboz Surgical Instrument, catalog number: RS-9262 )
  5. Wound clip remover 4’’ (Roboz Surgical Instrument, catalog number: RS-9263 )
  6. Ear punch; 2 mm; 2’’ length (Roboz Surgical Instrument, catalog number: 65-9900 )
  7. Micro dissecting scissors 4’’ straight sharp/sharp (Roboz Surgical Instrument, catalog number: RS-5912 )
  8. Moloney forceps; serrated; slight curve; 4.5’’ length (Roboz Surgical Instrument, catalog number: RS-8254 )
  9. Micro dissecting forceps; serrated, full curve, 4’’ length (Roboz Surgical Instrument, catalog number: RS-5137 )
  10. Isoflurane chamber and nose cone
  11. Fisher scientific slide warmer model 77 (Thermo Fisher Scientific, Fisher Scientific, catalog number: 12-594 )
  12. Oster Cord/Cordless trimmer (Patterson Veterinary Supply, catalog number: 07-880-1877 )
  13. Culture hood

Procedure

  1. Preparing cells for transplantation
    1. Grow and expand cells according to recommended conditions. If cells were thawed from cryogenic storage, make sure to passage cells at least two times before harvesting for transplantation.
    2. Harvest cells while in exponential growth phase (approximately 80-90% confluence) using trypsin or appropriate enzymes for the specific cell line. Suspend cells in medium plus serum.
    3. Count cells.
    4. Spin cells at 225 x g at 4 °C for 5 min. Resuspend cells in HBSS to a concentration of 0.5-2 million cells/200 µl.
    5. Place cells on ice and transport to vivarium.

Notes:

  1. Not all cell lines will grow in vivo. It is highly recommended that a pilot study is performed to determine the optimal concentration of cells needed to develop tumors that grow to 100 mm3 within 2-5 weeks from injection.
  2. Cells should be tested to ensure that they are not contaminated with viruses that are harmful to immunodeficient mice. This can be done using a traditional murine antibody production (MAP) test or one of the more sensitive PCR-based tests, such as the Research Animal Diagnostic Laboratory (RADIL) infectious microbe PCR amplification test (IMPACT) (IDEXX BioResearch).

  1. Subcutaneous injection or mammary fat pad injection of cells
    Note: Please view video for additional information on procedure (Video 1).

    Video 1. Preparation for xenograft procedures

    1. If using immunodeficient mice with hair, shaving with electric clippers may be necessary.
      1. For subcutaneous injections, shave the right, dorsal flank of the mouse from the hindlimb to the forelimb.
      2. For mammary fat pad injections, locate the fourth mammary fat pad (on the ventral side) and shave the fur over the fourth mammary fat pad and up around the dorsal side of the mouse.
    2. Fill 25 gauge needle with appropriate volume (typically 0.1-0.2 ml), carefully expelling any bubbles that may enter the syringe.
    3. Injecting the cells:
      1. For subcutaneous injections, restrain the animal in an upright position by grasping the skin over the shoulders with the thumb and index finger so that the fore legs are extended out to the side, keeping the front feet from pushing the syringe away.
        1. Gently insert the needle, bevel up, into the skin on the dorsal flank. Proceed to insert the needle slightly deeper than the skin to reach the subcutaneous pocket (deeper than the skin but avoid the underlying muscle layer).
        2. Take care to direct the needle away from the fingers that are restraining the mouse.
        3. Gently expel the contents of the syringe. The contents of the syringe should inject easily without resistance. 
        4. Resistance will indicate improper placement of the syringe, in either the muscle or skin. If in the muscle, a droplet of blood might be observed following the withdrawal of the syringe. If in the skin, the skin will expand and a whitish bleb (raised surface) will appear.
      2. For mammary fat pad injections, restrain the animal in an upright position by grasping the skin over the shoulders with the thumb and middle finger so that the fore legs are extended out to the side.
        1. Gently insert the needle into the 4th mammary fat pad. The needle should be inserted proximal to the nipple, bevel up, and 2-4 mm under the skin. In athymic nude mice you will be able to observe whether the needle is in the fat pad.
        2. Gently expel the contents of the syringe. The contents of the syringe should inject easily without resistance. 
        3. Resistance will indicate improper placement of the syringe, in either the muscle or skin. 
    4. At this point, the mouse may be placed back in the cage.

Note: To practice proper placement, dyes such as trypan blue can be injected into the subcutaneous pocket or the mammary fat pad. It is important to avoid injecting into the inguinal lymph node in the 4th mammary fat pad.

  1. Preparing primary tumor tissue for transplantation
    It is recommended that primary and serial tumor transplantation be performed with tumor fragments as opposed to single cell preparations in order to maintain the molecular and histological heterogeneity. 
    1. When receiving patient samples, always keep them on ice until the tissue is processed.
    2. Using a dedicated tissue culture hood for human tissues, take the tissue sample and put it onto a sterile 100 mm tissue culture dish.
    3. Pipette 1 ml PBS with 1% penicillin-streptomycin onto the tissue to keep it moist. This also helps to prevent the tissue from sticking to your instruments while processing.
    4. Cut the tumor into implantable pieces (approximately 3 x 3 x 3 mm) using sterile scissors and forceps.
    5. Place the tumor pieces in ice-cold PBS with 1% penicillin-streptomycin in either an Eppendorf tube or 15 ml conical tube depending on the volume of tumor pieces and put on ice.
    6. If needed, snap freeze the remaining tumor in liquid nitrogen for future analysis.
    7. Spray all instruments with 70% ethanol, then wash with surgical cleaner, and sterilize by autoclaving.
    8. Follow the surgical procedures below once the tumor pieces have been transported to the vivarium.

Note: Patient samples are not routinely tested for bloodborne pathogens. Proper technique and safety protocols should be followed when handling human tissue.

  1. Preparing cryopreserved tumor tissue for transplantation
    1. Thaw the tumor in the cryogenic vial in a 37 °C water bath for approximately 2 min.
    2. Using a dedicated tissue culture hood for human tissues, take out the tumor from the cryogenic vial with sterile forceps and place in a sterile 100 mm tissue culture plate.
    3. If the tumor tissue is larger than 27 mm3, cut the tumor into implantable size chunks (approximately 3 x 3 x 3 mm) using sterile scissors.
    4. Place the tumor pieces in ice-cold PBS with 1% penicillin-streptomycin in either an Eppendorf tube or 15 ml conical tube depending on the volume of tumor pieces and put on ice.
    5. Follow the surgical procedure below once the tumor pieces have been transported to the vivarium.

  2. Harvesting patient-derived xenografts for serial transplantation
    Note: Please view video for additional information on procedure (Video 2).

    Video 2. Harvesting patient-derived xenografts for serial transplantation

    For serial transplantation or transplantation for drug studies, it is often possible to obtain 20-50 transplantable pieces from a single tumorgraft. It is critical that necrotic regions of the tumor are avoided.
    1. Euthanize the mouse carrying the patient-derived tumor according to your institutional IACUC protocol. Ideally euthanasia should be performed in the same room as the tumor implantation surgery.
    2. Immediately following euthanasia, remove the PDX tumor using aseptic technique as follows:
      1. Place the mouse on an absorbent underpad and spray the entire mouse down with 70% ethanol, concentrating on the tumor area.
      2. Set up two sets of sterile forceps and scissors and lay them in 100 mm culture dishes to avoid surface contamination.
      3. Remove the tumor using the two separate sets of forceps and scissors. The first set is used to cut into the skin (dirty side). The second set should be used for internal dissection (clean side) of the tumor. Be careful not to let the tumor touch the outside skin of the mouse or any other sites of potential contamination. 
      4. Place the tumor in a new sterile 100 mm tissue culture dish.
    3. Rinse the tumor with ice-cold PBS with 1% penicillin-streptomycin (approximately 1 ml per tumor is sufficient).
    4. Cut and process any tissue needed for cryopreservation, histology, or snap freezing before cutting pieces for transplant.
    5. Cut the tumor into as many implantable pieces as needed (approximately 3 x 3 x 3 mm), working as quickly as possible.
    6. Place the pieces in ice-cold PBS with 1% penicillin-streptomycin in a new sterile culture dish on ice.
    7. Proceed to transplanting the tumor pieces into new mice as described in Procedure F.
    8. Dispose of the donor mouse carcass.

  3. Mammary fat pad or subcutaneous transplantation of tumorgrafts
    Note: Please view video for additional information on procedure (Video 3).

    Video 3. Mammary fat pad transplantation of tumorgrafts

    1. Tape a sterile absorbent underpad to the benchtop (30 x 15 cm) for surgical procedures.
    2. Anesthetize the mice for transplant in batches of five or as a cage group to minimize the length of time they are under anesthesia. Described below is the procedure for using the isoflurane as an anesthesia at our institution.
      1. Tape the isoflurane nosecone to the left edge of the absorbent underpad.
      2. Check the waste gas scavenging system on the anesthesia machine before work begins. Open the valve to the house vacuum to allow negative pressure to be maintained in the gas scavenging interface device.
      3. Check the position of the valve leading to the nose cone to make sure it is in the ‘off’ position. Failure to monitor the position of the valves when beginning work and throughout the procedure may allow excess gas to escape into the lab space and cause a health danger to personnel in lab area.
      4. Open the valves to the oxygen tank. Adjust the pressure to between 10-15 psi and the flow to 1 L per minute. Note that the flow may need to be increased, depending on the number of mice that are being anesthetized.
      5. Open the anesthesia chamber and place the mouse inside. Close the lid to the chamber before starting the anesthesia.
      6. Adjust the vaporizer to 3 percent isoflurane in oxygen.
      7. Monitor the animal. When cessation of movement occurs, close the valve to the chamber and open the valve to the nose cone. If more than one mouse is being anesthetized, maintain the anesthesia flow to the chamber.
    3. While mice are being anesthetized, set up the rest of the surgery station keeping everything as aseptic as possible (open sterile scissors, forceps, and a trocar syringe and set on the inside of the sterilization pouch or into a sterile 100 mm tissue culture dish, open the betadine swabs, set out sterile alcohol pads, and load staples into the surgical stapler).
    4. Remove one mouse from the isoflurane chamber after it has reached surgical anesthesia level.
    5. Pull up the skin at the base of the mouse’s neck and inject 100 µl of 5 mg/kg ketoprofen (or equivalent analgesic) subcutaneously with a 28 gauge needle. 
    6. For mammary fat pad transplantation:
      1. Open valve to nose cone and place the mouse’s nose in the nosecone with the dorsal side of the mouse facing down toward the absorbent underpad.
      2. Locate the fourth mammary fat pad and shave the fur over the fourth mammary fat pad and up around the dorsal side of the mouse using electric clippers.
      3. Swab the shaved skin with betadine.
      4. Wipe off the betadine with sterile alcohol prep pads. 
      5. Pull up the skin right above the nipple with forceps and cut a small incision using scissors (4-5 mm) horizontally across the fourth mammary fat pad.
      6. Use the trocar syringe to make a vertical guide path underneath the skin for the tumor, following the fat pad around to the back of the mouse.
      7. Remove the trocar and place a tumor piece into the tip of the syringe with forceps.
      8. Transplant the tumor piece into the fourth mammary fat pad by inserting the trocar back into the incision and pushing it around to the back of the mouse through the guide path, expelling the tumor.
      Note: The farther the tumor engrafts away from the nipple or groin area of the mouse, the easier it will be to measure with calipers.
    1. Alternatively, a pair of small, curved forceps may be used in place of a trocar to make a guide path under the skin for the tumor. Gently insert the forceps into the fourth mammary fat pad and move around in a small circular motion to create a pocket for the tumor. These same forceps will then place the tumor into the fourth mammary fat pad.
    2. Slowly and gently remove the trocar, ensuring that the tumor remains deep within the fourth mammary fat pad.
    1. For subcutaneous transplantation:
      1. Open valve to nose cone and place the mouse’s nose in the nosecone with the ventral side of the mouse facing down toward the absorbent underpad.
      2. Shave the right, dorsal flank of the mouse from the hindlimb to the forelimb.
      3. Swab the shaved skin with betadine.
      4. Wipe off the betadine with sterile alcohol prep pads. 
      5. Pull up the skin right on the dorsal flank with forceps and cut a small incision using scissors (4-5 mm) horizontally.
      6. Use the sterile scissors or forceps to make a small pocket under the skin and place a tumor fragment under the skin. Make sure to place the tumor fragment at least 3 mm from the opening.
    2. Grasp both ends of the initial surgical opening with forceps, pull up, and adjust your hands and the forceps so that you are grasping the edges of the incision with one forcep. While still pulling up the skin, staple the incision shut. Avoid stapling into the body cavity of the mouse.
    3. Transfer the mouse to a temporary, clean cage on a 37 °C warmer. Monitor the animal closely until it regains consciousness.
    4. Proceed with the tumor implants on the remaining four mice, transferring each to the warming cage after surgery. When all mice have regained consciousness and are moving normally, they can be transferred to their standard housing cage. 
    5. When surgery has been completed on the fifth mouse, you may transfer five additional mice to the isoflurane chamber to anesthetize.
    6. When finished, turn off the isoflurane machine and spray all surfaces with 70% ethanol and wipe down, followed by antiseptic, such as Clidox.
    7. Clean all instruments by rinsing in water, then washing with surgical instrument cleaner. Autoclave to sterilize.
    8. Remove staples after 7-14 days.

  4. Implantation of estrogen pellets
    Note: Please view video for additional information on procedure (Video 4).

    Video 4. Subcutaneous transplantation of tumorgrafts

    If using breast cancer cells or a patient-derived breast cancer tumor that is estrogen dependent, it is necessary to supplement tumor growth with estrogen. Estrogen pellets that consistently release over 60-90 days are recommended.
    1. This procedure requires anesthesia and may be performed before or after tumorgraft implantation or cell inoculation.
    2. Anesthetize the mouse with isoflurane according to your institutional IACUC protocol.
    3. Shave the hair off the dorsal side of the neck above the shoulder blades. The use of depilatory cream is not recommended with immunodeficient models with a SCID background.
    4. Swab the exposed skin with a betadine swab and wipe off with sterile alcohol pad.
    5. Make an incision equal in diameter to that of the pellet.
    6. Pull up the skin on one side of the incision using forceps and make a pocket horizontally with a pair of forceps or scissors about 1 cm beyond the incision site.
    7. Insert the estrogen pellet into the pocket using forceps.
    8. Staple or glue the incision shut (some mice will not tolerate a staple in their neck and will scratch it out, leading to loss of the estrogen pellet).
    9. Remove staple after 7-14 days.

  5. Cryopreservation of tumor tissue
    1. Prepare freezing medium: Media + 10% FBS + sterile 10% DMSO.
    2. Place aseptically harvested tumor tissue into a sterile culture dish and rinse with approximately 5 ml of sterile PBS containing 1% penicillin-streptomycin.
    3. Visibly examine the tissue to carefully remove and discard any necrotic and non-tumor tissue from the tumor specimen.
    4. Using forceps and scalpel, dissect the tumor tissue into 3 x 3 x 3 mm fragments; depending on the tumor type, some tissue may be more easily dissected by teasing apart with forceps rather than by cutting with a scalpel.
    5. Label cryogenic vials with tumor tissue ID and date harvested and add 1 ml freezing medium.
    6. Place a single tumor fragment into a cryogenic vial, tighten cap and place in the slow-rate freezing apparatus.
    7. Replace lid on freezing apparatus and place into -80 °C freezer overnight. The tumor tissue will freeze at a rate of 0.5 °C/min to -20 °C and 1 °C/min to -80 °C.
    8. Following overnight freezing, store cryogenic vials in a liquid nitrogen freezer and record location and date of cryostorage.

  6. Tumor measurement and drug treatment (oral gavage and intraperitoneal injection)
    1. Tumor growth should be measured 2 to 3 times weekly using calipers (length, width, and height).
    2. Gently restrain the mouse in a manner that allows access to the tumor.
    3. Bring the calipers to the longest side of the tumor to measure the length. Gently squeeze the calipers together until the borders of the tumor are within the caliper jaws.
    4. Bring the calipers to the narrower side of the tumor to measure the width.
    5. The height measurement may be taken by trying to use the calipers to gauge the height of the tumor. Alternatively, a plastic ruler may be placed flush with the base of the tumor and the height of the tumor read.
    6. Record length, width, and height measurements appropriately.
      Note: For most consistent results, the same person should measure tumors for the duration of the study. Otherwise, variability may be observed since each person may squeeze the tumors differently during measurement.
    7. Tumor volume can be calculated by an ellipsoid volume formula (π/6 x L x W x H) or the volume formula (L x W x H) (Tomayko and Reynolds, 1989).
    8. The tumor volume at which drug treatment begins is dependent on the goal of the experiment, but in general the tumor should be at least 100 mm3 to ensure established tumor growth. If the goal of the experiment is to test ‘prevention’ of tumor growth, the drug treatment timeline should be adjusted.
      1. For experiments conducted on established tumors, an approximate tumor volume average of 150 mm3 is used as the drug treatment starting size. When a cohort of animals has an average tumor size of 150 mm3, the mice are randomized into treatment groups. Animals with tumors less than 115 mm3 are not included in the first cohort and will be randomized into a second cohort of mice and enrolled on study at a later date.
        Note: The rate and uniformity of tumor growth is entirely dependent on the properties of the tumor cell line or tumorgraft. In some experiments, all the transplanted mice will be able to start treatment at the same time and in some experiments, it may be necessary to divide the animals into several cohorts that are enrolled on study with at various dates.
      2. After randomization the mice are treated with vehicle or drug depending on their assigned treatment group.
      3. The dosage, dosing schedule (weekly, daily, twice daily, etc.), and vehicle is determined by the chemical compound. This information is often available in the published literature or from the pharmaceutical company that provides the compound.
    9. Oral gavage dosing.
      1. Weigh the animals to determine the average weight and dose/volume needed. The maximum volume that can be gavaged in a mouse is 200 µl.  
      2. Prepare the 22 gauge gavage needle by attaching to a 1 cc syringe.
      3. Restrain the animal in an upright position by grasping the skin over the shoulders with the thumb and index finger so that the fore legs are extended out to the side, keeping the front feet from pushing the gavage tube away.
      4. Hold the animal’s head in place by gently extending the head back (this extension of the head creates a straight line through the neck and esophagus).
      5. The gavage needle is used to measure the distance from the mouth to the last rib (stomach) of the mouse. This is approximately how far the gavage needle must be inserted to ensure an effective procedure.
      6. Insert the bulb of the needle into the corner of the mouth. Reposition the bulb toward the center and run it along the roof of the mouth.
      7. Using the needle as a lever, gently push back (upright) the head and the nose of the mouse. (The restraining hand is kept upright, not bent back.) 
      8. Slowly feed the gavage needle down the esophagus to the pre-measured distance. If resistance is felt, do not advance the needle any further. If necessary, retract the needle until the bulb is at the front of the mouth (but not out) and reinsert. Do not attempt insertion more than 3 times. If insertion fails after 3 attempts the animal must be allowed to rest for 2 h. After the rest period, the insertion may be attempted an additional 3 times. If the second round fails, the attending veterinarian should be consulted before continuing any oral gavage.
      9. Advance the needle, extending the head and the nose further back or upright.
      10. Make sure the needle is fully inserted into the pre-measured target point prior to pushing the plunger.
      11. Push the plunger to release the contents into the animal’s stomach.
      Note: Gavage needles may be used on multiple animals within a study. However, needles should be changed between treatments.
    10. Intraperitoneal (IP) injection
      1. To locate the point of entry for the needle draw an imaginary line across the abdomen just above the knees. The needle will be inserted along this line on the animal’s right side and close to the midline. In female animals, the point of entry is cranial to and slightly medial of the last nipple. Inserting the needle on the mouse’s right side avoids the cecum.
      2. To restrain the mouse: grasp the skin over the shoulders with the thumb and index finger. Tilt the head so that it is facing downward and the abdomen is exposed. To perform an IP injection, the mouse must be well restrained so that it cannot move during the procedure. This avoids traumatizing the organs once the needle has entered the abdomen.
      3. Insert the 27 gauge needle into the abdomen at about a 30-degree angle. The shaft of the needle should enter to a depth of about half a centimeter.
      4. Prior to injection, aspirate to be sure that the needle has not penetrated a blood vessel, the intestines, or the urinary bladder.
        1. Greenish brown aspirate indicates needle penetration into intestines.
        2. Yellow aspirate indicates needle penetration into the bladder.
        3. If any fluid is aspirated, your solution is contaminated and must be discarded and the procedure repeated with a new syringe and needle.
        4. If no fluid is aspirated, you may inject.
      5. Withdraw the needle and return the mouse to its cage.

Data analysis

For drug treatment experiments, mice are randomized into control or treatment groups based on tumor size. The number of mice needed for each experiment/treatment group is determined by a power analysis. Tumor volume is longitudinally monitored pre- and post-treatment. Biostatisticians are blinded to treatment status to mitigate bias being introduced to the statistical analysis. Linear mixed-effects models are used test for significant differences in drug response across treatment arms and the XenoCat modeling framework is leveraged to increase statistical power if poorly growing xenograft/PDX subjects are present (Laajala et al., 2012). Appropriate model contrasts are invoked to formally test for synergism or antagonism in drug response. Bonferroni corrections are applied to control the familywise error rate.

Notes

  1. Before running a large xenograft study with cell lines, it is recommended that a small pilot study is run to determine the tumor take rate. This will help determine the number of additional mice that are needed if the take rate is less than 100%.
  2. The success and viability of cell line xenografts can be affected by the confluency at the time of harvesting. As noted above it is best to harvest when cells are in the exponential growth stage. If the tumor viability is poor with a cell line xenograft, Matrigel can also be added to the cell-media injection mixture in order to provide an enriched stromal environment for engraftment.
  3. With patient-derived xenograft models, engraftment is highly dependent on the tumor type. Metastatic breast cancer can have a take rate of 25%, yet treatment-naïve primary breast cancer has a take rate close to 5% in our studies.

Acknowledgments

We would like to thank Bryn Eagleson and the VARI Vivarium staff for their expertise. The protocols described here were used in multiple studies supported by The Breast Cancer Research Foundation, Muskegon Tempting Tables, and the Van Andel Foundation.

References

  1. Bosma, M. J. and Carroll, A. M. (1991). The SCID mouse mutant: definition, characterization, and potential uses. Annu Rev Immunol 9: 323-350.
  2. DeRose, Y. S., Gligorich, K. M., Wang, G., Georgelas, A., Bowman, P., Courdy, S. J., Welm, A. L. and Welm, B. E. (2013). Patient-derived models of human breast cancer: protocols for in vitro and in vivo applications in tumor biology and translational medicine. Curr Protoc Pharmacol Chapter 14: Unit14 23.
  3. DeRose, Y. S., Wang, G., Lin, Y. C., Bernard, P. S., Buys, S. S., Ebbert, M. T., Factor, R., Matsen, C., Milash, B. A., Nelson, E., Neumayer, L., Randall, R. L., Stijleman, I. J., Welm, B. E. and Welm, A. L. (2011). Tumor grafts derived from women with breast cancer authentically reflect tumor pathology, growth, metastasis and disease outcomes. Nat Med 17(11): 1514-1520.
  4. Flanagan, S. P. (1966). ‘Nude’, a new hairless gene with pleiotropic effects in the mouse. Genet Res 8(03): 295-309.
  5. Fruchter, R. G., Nayeri, K., Remy, J. C., Wright, C., Feldman, J. G., Boyce, J. G. and Burnett, W. S. (1990). Cervix and breast cancer incidence in immigrant Caribbean women. Am J Public Health 80(6): 722-72
  6. Gao, H., Korn, J. M., Ferretti, S., Monahan, J. E., Wang, Y., Singh, M., Zhao, C., Schnell, C., Yang, G., Zhang, Y., Balbin O. A., Barbe, S., Cai, H., Casey, F., Chatterjee, S., Chiang D. Y., Shannon, C., Cogan, S. M., Collins, S. D., Dammassa, E., Ebel, N., Embry, M., Green, J., Kauffmann, A., Kowal, C., Leary, R. J., Lehar, J., Liang, Y., Loo, A., Lorenzana, E., McDonald III, E. R., McLaughlin, M. E., Merkin, J., Meyer, R., Naylor, T. L., Patawaran, M., Reddy, A., Röelli, C., Ruddy, D. A., Salangsang, F., Santacroce, F., Singh, A. P., Tang, Y., Tinetto, W., Tobler, S., Velazquez, R., Venkatesan, K., Arx, F. V., Wang, Q., Wang, Z., Wiesmann, M., Wyss, D., Xu, F., Bitter, H., Atadja, P., Lees, E., Hofmann, F., Li, E., Keen, N., Cozens, R., Jensen, M. R., Pryer, N. K., Williams, J. A. and Sellers, W. R., (2015). High-throughput screening using patient-derived tumor xenografts to predict clinical trial drug response. Nat Med 21(11): 1318-1325.
  7. Hait, W. N. (2010). Anticancer drug development: the grand challenges. Nat Rev Drug Discov 9(4): 253-254.
  8. Laajala, T. D., Corander, J., Saarinen, N. M., Makela, K., Savolainen, S., Suominen, M. I., Alhoniemi, E., Makela, S., Poutanen, M. and Aittokallio, T. (2012). Improved statistical modeling of tumor growth and treatment effect in preclinical animal studies with highly heterogeneous responses in vivo. Clin Cancer Res 18(16): 4385-4396.
  9. Prochazka, M., Gaskins, H. R., Shultz, L. D. and Leiter, E. H. (1992). The nonobese diabetic scid mouse: model for spontaneous thymomagenesis associated with immunodeficiency. Proc Natl Acad Sci U S A 89(8): 3290-3294.
  10. Sausville, E. A. and Burger, A. M. (2006). Contributions of human tumor xenografts to anticancer drug development. Cancer Res 66(7): 3351-3354, discussion 3354.
  11. Shinkai, Y., Rathbun, G., Lam, K. P., Oltz, E. M., Stewart, V., Mendelsohn, M., Charron, J., Datta, M., Young, F., Stall, A. M. and et al. (1992). RAG-2-deficient mice lack mature lymphocytes owing to inability to initiate V(D)J rearrangement. Cell 68(5): 855-867.
  12. Shultz, L. D., Lyons, B. L., Burzenski, L. M., Gott, B., Chen, X., Chaleff, S., Kotb, M., Gillies, S. D., King, M., Mangada, J., Greiner, D. L. and Handgretinger, R. (2005). Human lymphoid and myeloid cell development in NOD/LtSz-scid IL2R γnull mice engrafted with mobilized human hemopoietic stem cells. J Immunol 174(10): 6477-6489.
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简介

衍生自人类癌细胞的体内异种移植模型已经成为评估癌症遗传学驱动因素的黄金标准,并且是用于评估癌症治疗药物功效的有价值的临床前模型。最近,已经开发了来自多种肿瘤类型的患者衍生的移植物,并且显示出更准确地概括了癌症的分子和组织学异质性。在这里,我们详细介绍了从乳腺癌组织,基于细胞的异种移植模型,连续肿瘤移植,肿瘤测量和药物治疗开发患者衍生异种移植模型的程序。

背景 异种移植模型已经成为研究癌症基因驱动因素和确定癌症治疗药物的潜在功效的有力方法。发现T细胞缺乏的无胸腺裸鼠(nu / nu)小鼠和T细胞和B细胞缺陷型严重联合免疫缺陷(scid / scid)小鼠(Flanagan, 1966; Bosma和Carroll,1991)。由于这些发现,其他免疫受损的小鼠模型已经变得可用,包括重组激活基因2(Rag2)敲除小鼠,非肥胖(NOD) - 小鼠和NOD-scid < IL2Rgamma(null)小鼠(也称为NSG小鼠)(Shinkai等人,1992; Prochazka等人,1992; Shultz等人,2005)。这些免疫受损的小鼠模型已经开发出许多和多样化的人类癌症体内模型。
&nbsp;在开发包括注射或植入部位的异种移植模型时,应考虑几个选项。由于生长测量和成像的肿瘤可及性,皮下异种移植物通常用于体内研究;然而,该模型的显着限制是大多数癌细胞缺乏正常的基质微环境。原位异种移植物提供互补的基质微环境;然而,根据原位点,包括更复杂的外科手术,测量肿瘤生长或反应的难度以及啮齿动物基质的限制(Talmadge等人,2007),该途径也存在缺点。许多癌症研究尤其是乳腺癌研究中广泛应用原位异种移植物。注入乳腺脂肪垫是一个相对简单的程序,允许乳腺癌细胞的可见和可测量的生长。尽管乳腺脂肪垫为乳腺癌细胞提供了互补的组织部位,但重要的是注意到人和啮齿动物乳腺基质和激素环境之间存在明显差异。
&nbsp;异种移植模型被广泛用作癌症治疗功效的预测模型。对于临床前研究,必须在体内评价药物功效和潜在的毒性。尽管在体内临床前研究是有价值的,但结果并不一致地转化为临床,并且对这些研究的意义进行了辩论(Talmadge等人,2007; Sausville和Burger,2006)。在设计药物研究(如合适的细胞系(或PDX模型)),剂量和给药方案以及统计分析时,需要考虑几个变量。应仔细考虑这些因素,以最为模仿人类癌症进展和治疗反应。
&nbsp;最近在患者来源的肿瘤异种移植物(PDX)的发展方面取得了重大进展。 PDX模型具有保持原始肿瘤的分子和组织学异质性的优点(DeRose等人,2011)。此外,与标准细胞培养异种移植模型相比,它们已被证明在预测药物反应方面是优越的(Hait,2010; Fruchter等,1990; Voskoglou-Nomikos等人,2003; Gao等人,2015)。最近的研究已经提高了建立乳腺癌PDX模型的成功,该模型概括了存在于乳腺癌中的分子,基质和表型异质性(DeRose等人,2013)。总体而言,基于细胞系和患者来源的异种移植物是研究癌症起始,进展和治疗反应的必要模型。在这里,我们描述了开发基于细胞的异种移植模型,来自乳腺癌组织的患者衍生异种移植模型,连续肿瘤移植,肿瘤测量和药物治疗的方案。

关键字:患者源性异种移植瘤, 肿瘤移植, 乳房脂肪垫, 肿瘤测量, 剂量

材料和试剂

  1. 25号针(用于接种)(BD,目录号:305127)
  2. 组织培养处理培养皿(Corning,目录号:430293)
  3. Eppendorf管(1.5ml)(Eppendorf,目录号:022363204)
  4. Falcon 15 ml锥形离心管(Corning,目录号:352196)
  5. 低温小瓶
  6. 套管针注射器,10号(美国创新研究,目录号:MP-182)
  7. 1cc U-100胰岛素注射器,28磅x½"针(用于酮洛芬注射)(BD,目录号:329410)
  8. 口服管饲技巧,22磅x 1",1¼毫米球(Cadence,目录号:7901)
  9. 结核菌素注射器,1ml(用于接种)(BD,目录号:309659)
  10. Betadine溶液拭子棒(Thermo Fisher Scientific,Fisher Scientific,目录号:19-061617)
  11. 无菌酒精制品垫,70%异丙醇(Thermo Fisher Scientific,Fisher Scientific,目录号:22-363-750)
  12. 无菌细胞制剂或患者来源的肿瘤组织
  13. 4-7周龄的Athymic裸鼠[Crl:NU(NCr)-Foxn1 nu ](Charles River Laboratories Internetional,目录号:490)
  14. 4-7周龄NSG小鼠(NOD γ,NOD-scid IL2Rg null ,NOD- scid IL2Rgamma null )(JACKSON LABORATORY,目录号:005557)
  15. 胰蛋白酶或适当的酶
  16. 汉克斯平衡盐溶液(HBSS)(Thermo Fisher Scientific,Gibco TM,目录号:14170112)
  17. 研究动物诊断实验室(RADIL)感染性微生物PCR扩增试验(IMPACT)(IDEXX BioResearch)
  18. 磷酸盐缓冲盐水(PBS)pH 7.4(Thermo Fisher Scientific,Gibco TM,目录号:10010049)
  19. 青霉素 - 链霉素(5,000U/ml)(Thermo Fisher Scientific,Gibco TM,目录号:15070063)
  20. 70%乙醇
  21. 手术清洁剂
  22. 异氟烷液体吸入剂OD 99.9%(Henry Schein Medical,目录号:1182097)
  23. Ketofen ®(酮洛芬)(Patterson Veterinary Supply,目录号:10004029)
  24. 吸收性垫片(Thermo Fisher Scientific,Fisher Scientific,目录号:S67011)
  25. 10%中性福尔马林(Sigma-Aldrich,目录号:HT501128-4L)
  26. FBS
  27. DMSO
  28. 可选材料和试剂
    1. 17β-雌二醇,60天释放丸(美国创新研究,目录号:SE-121)
    2. 兽医级手术胶(Patterson兽医用品,目录号:07-805-5031)
    3. 低温培养基(见下表)
    4. 细胞冷冻容器(Corning,目录号:432010)
    5. 内螺纹低温小瓶,2.0毫升(康宁,目录号:430488)
    6. 液氮

设备

  1. 数字卡钳(Thermo Fisher Scientific,Fisher Scientific,目录号:06-664-16或美国三丰,目录号:500-163-30)
  2. 安全解剖刀(Merit Medical Systems,目录号:SMS210)
  3. 伤口夹钳9毫米(Roboz手术器械,目录号:RS-9260)
  4. 9 mm伤口夹(Roboz Surgical Instrument,目录号:RS-9262)
  5. 伤口夹去除器4"(Roboz手术器械,目录号:RS-9263)
  6. 耳冲2毫米; 2"长(Roboz手术器械,目录号:65-9900)
  7. 微解剖剪刀4''锋利尖锐(Roboz手术器械,目录号:RS-5912)
  8. 莫洛尼钳锯齿状轻微曲线4.5"长(Roboz手术器械,目录号:RS-8254)
  9. 微解剖钳锯齿状,全曲线,4"长(Roboz手术器械,目录号:RS-5137)
  10. 异氟醚室和鼻锥
  11. Fisher科学幻灯片加热器型号77(Thermo Fisher Scientific,Fisher Scientific,目录号:12-594)
  12. Oster Cord/Cordless trimmer(Patterson Veterinary Supply,目录号:07-880-1877)
  13. 文化罩

程序

  1. 准备移植细胞
    1. 根据推荐条件生长和扩大细胞。如果细胞从低温储存中解冻,在收获前应确保至少两次传代细胞进行移植。
    2. 在使用胰蛋白酶或特定细胞系合适的酶的指数生长期(约80-90%汇合)收获细胞。将细胞悬浮于培养基加血清。
    3. 计数单元格。
    4. 旋转细胞在225℃下在4℃下5分钟。将HBSS中的细胞重悬于0.5-2百万个细胞/200μl的浓度。
    5. 将细胞放在冰上并运送至生殖细胞。

注意:

  1. 不是所有的细胞系都将在体内生长。强烈建议进行一项初步研究,以确定在注射后2-5周内发展成100 mm 3/sup的肿瘤所需的细胞的最佳浓度。
  2. 细胞应该进行测试,以确保它们不会被对免疫缺陷小鼠有害的病毒污染。这可以使用传统的鼠抗体生产(MAP)测试或更灵敏的基于PCR的测试之一进行,例如研究动物诊断实验室(RADIL)感染性微生物PCR扩增试验(IMPACT)(IDEXX BioResearch)。 em>

  1. 皮下注射或乳腺脂肪垫注射细胞
    注意:请查看视频以了解有关步骤的其他信息(视频1)。

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    视频1.异种移植手术的准备
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    1. 如果使用带头发的免疫缺陷小鼠,可能需要用电动剪刀剃须。
      1. 对于皮下注射,将后腿的右侧背侧剃刮至前肢。
      2. 对于乳房脂肪垫注射,定位第四个乳腺脂肪垫(在腹侧),并将毛皮刮去第四个乳腺脂肪垫,并向上靠近鼠标的背侧。
    2. 以适当体积(通常为0.1-0.2毫升)填充25号针头,仔细排出可能进入注射器的任何气泡。
    3. 注入细胞:
      1. 对于皮下注射,通过用拇指和食指抓住肩膀上的皮肤使得前腿伸出到侧面,保持前脚不要将注射器推开,从而将动物限制在直立位置。
        1. 轻轻地将针头倒入背部的皮肤上。继续将针头插入比皮肤稍深的位置,以便到达皮下的口袋(比皮肤深,但避免下面的肌肉层)。
        2. 注意将针头指向限制鼠标的手指。
        3. 轻轻排出注射器的内容物。注射器的内容物容易注射,无阻力。
        4. 阻力将表示注射器在肌肉或皮肤中的不正确放置。如果在肌肉中,注射器撤出后可能会观察到一滴血迹。如果在皮肤中,皮肤会膨胀,会出现发白的泡沫(凸起的表面)。
      2. 对于乳房脂肪垫注射,通过用拇指和中指抓住肩膀上的皮肤使得前腿伸出到侧面,使动物处于直立位置。
        1. 轻轻地将针插入到第4个乳房脂肪垫。针头应插入乳头的近侧,向上倾斜,皮肤下方为2-4毫米。在无胸腺裸鼠中,您可以观察针是否在脂肪垫中。
        2. 轻轻排出注射器的内容物。注射器的内容物容易注射,无阻力。
        3. 阻力将表示注射器在肌肉或皮肤中的不正确放置。
    4. 此时,鼠标可以放回笼子中。

注意:要进行适当的放置,染色如台盼蓝可以注射到皮下袋或乳房脂肪垫中。重要的是避免注射到乳房脂肪垫的腹股沟淋巴结中。

  1. 准备原发肿瘤组织进行移植
    建议使用肿瘤片段进行原代和连续的肿瘤移植,而不是单细胞制剂,以保持分子和组织学异质性。
    1. 收到患者样本时,请始终将其保持在冰上,直至组织被处理
    2. 使用人体组织的专用组织培养罩,取组织样品并放入无菌的100毫米组织培养皿中。
    3. 将1ml含1%青霉素 - 链霉素的PBS吸移到组织上以保持其湿润。这也有助于防止组织在处理时粘到仪器上。
    4. 使用无菌剪刀和镊子将肿瘤切割成可植入的块(约3 x 3 x 3 mm)。
    5. 将肿瘤片放在含有1%青霉素 - 链霉素的冰冷的PBS中,根据肿瘤块的体积将其放入Eppendorf管或15ml锥形管中,并放在冰上。
    6. 如果需要,将剩余的肿瘤冻结在液氮中,以备将来分析
    7. 用70%乙醇喷洒所有仪器,然后用手术清洁剂清洗,并通过高压灭菌消毒
    8. 一旦肿瘤块被运送到vivarium,请按照下面的手术步骤。

注意:患者样品不是常规测试血源性病原体。处理人体组织时应遵循适当的技术和安全方案

  1. 准备冷冻保存的肿瘤组织进行移植
    1. 在37℃水浴中将低温小瓶中的肿瘤解冻约2分钟
    2. 使用人体组织的专用组织培养罩,用无菌镊子从低温小瓶中取出肿瘤,并置于无菌的100毫米组织培养板中。
    3. 如果肿瘤组织大于27 mm 3,使用无菌剪刀将肿瘤切割成可植入的大小块(约3 x 3 x 3 mm)。
    4. 将肿瘤片放在含有1%青霉素 - 链霉素的冰冷的PBS中,根据肿瘤块的体积将其放入Eppendorf管或15ml锥形管中,并放在冰上。
    5. 将肿瘤片段运送到生殖器后,按照下面的手术方法进行。

  2. 收集患者来源的异种移植物进行连续移植
    注意:请查看视频以了解有关步骤的其他信息(视频2)。

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    视频2.收集患者来源的异种移植物进行连续移植
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    对于用于药物研究的连续移植或移植,通常可以从单个肿瘤移植物获得20-50个可移植片。至关重要的是要避免肿瘤的坏死区域
    1. 根据您的机构IACUC方案对携带病人来源的肿瘤的小鼠进行安乐死。理想情况下,安乐死应在与肿瘤植入手术相同的房间内进行
    2. 安乐死后立即用无菌技术清除PDX肿瘤,如下所示:
      1. 将鼠标放在吸收垫上,用70%乙醇将整只小鼠向下喷,并集中在肿瘤区域上
      2. 设置两套无菌镊子和剪刀,放在100毫米培养皿中,以避免表面污染。
      3. 使用两套分开的镊子和剪刀去除肿瘤。第一套用于切入皮肤(脏面)。第二组应用于肿瘤的内部解剖(干净侧)。小心不要让肿瘤接触鼠标的外部皮肤或任何其他潜在污染的地方。 
      4. 将肿瘤置于新的无菌100毫米组织培养皿中。
    3. 用含有1%青霉素 - 链霉素的冰冷的PBS冲洗肿瘤(每个肿瘤约1ml)。
    4. 切割和处理切片之前冷冻保存,组织学或快速冷冻所需的任何组织
    5. 根据需要将肿瘤切割成尽可能多的植入物(约3 x 3 x 3 mm),尽可能快地工作。
    6. 将片放在含有1%青霉素 - 链霉素的冰冷的PBS中,放入冰上的新的无菌培养皿中。
    7. 继续将肿瘤块移植到新的小鼠中,如方法F所述
    8. 处置供体小鼠尸体。

  3. 乳腺脂肪垫或皮下移植肿瘤移植物
    注意:请查看视频以了解有关步骤的其他信息(视频3)。

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    视频3.乳腺脂肪移植肿瘤移植
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    1. 将无菌吸收垫放在台面上(30 x 15厘米)进行外科手术。
    2. 麻醉小鼠移植5批或笼子组,以尽量减少麻醉时间。下面描述的是在我们的机构使用异氟烷作为麻醉的程序。
      1. 将异氟烷胶囊粘贴到吸收垫的左边缘。
      2. 在工作开始前检查麻醉机上的废气清除系统。将阀门打开到室内真空,以便在气体清除界面装置中保持负压。
      3. 检查通向鼻锥的阀门的位置,以确保其处于"关闭"位置。在开始工作和整个程序过程中,不能监控阀的位置,可能会导致过量的气体逸出到实验室空间,对实验室的人员造成健康危害。
      4. 打开阀门到氧气罐。将压力调节至10-15 psi,流量每分钟达到1 L。请注意,根据正在麻醉的小鼠数量,流量可能需要增加。
      5. 打开麻醉室,将鼠标放在里面。在开始麻醉之前关闭房间的盖子。
      6. 将蒸发器调节至3%异氟醚在氧气中。
      7. 监视动物。当运动停止时,关闭阀门并打开阀门到鼻锥。如果不止一只老鼠被麻醉,请保持麻醉流向腔室。
    3. 当小鼠被麻醉时,建立手术台的其余部分,尽可能保持一切无菌(开放的无菌剪刀,镊子和套针管注射器,并置于灭菌袋的内部或放入无菌的100毫米组织培养皿中,打开betadine拭子,放出无菌酒精垫,并将钉书钉装入外科缝合器)。
    4. 达到手术麻醉水平后,从异氟醚室取出一只小鼠
    5. 拉起小鼠颈部的皮肤,并用28号针头皮下注射100μl5 mg/kg酮洛芬(或同等止痛剂)。
    6. 乳腺脂肪移植:
      1. 打开阀门到鼻锥,将鼠标的鼻子放在鼻锥上,鼠标的背面向下朝向吸收垫。
      2. 找到第四个乳腺脂肪垫,然后使用电动剪刀将皮毛剃除在第四个乳房脂肪垫上,并在鼠标的背侧上方。
      3. 用betadine擦拭剃光皮肤。
      4. 用无菌酒精制剂垫擦去betadine。
      5. 用镊子将乳头上方的皮肤拉起,并用剪刀(4-5毫米)横切第四个乳房脂肪垫切割小切口。
      6. 使用套管针注射器在肿瘤的皮肤下面形成一个垂直的引导路径,沿着靠近鼠标背部的脂肪垫。
      7. 取出套管针,用镊子将肿瘤块放入注射器的尖端。
      8. 通过将套管针插入切口并将其推向鼠标的后部通过引导路径将肿瘤块移植到第四乳腺脂肪垫中,排除肿瘤。
      注意:肿瘤移植距离鼠标的乳头或腹股沟区域越远,用卡尺测量越容易。
    1. 或者,可以使用一对小的弯曲的镊子来代替套管针,以在肿瘤的皮肤下面形成引导路径。轻轻地将镊子插入第四个乳房脂肪垫,并以小的圆周运动移动,以创建肿瘤的口袋。然后将这些相同的镊子放入第四个乳腺脂肪垫。
    2. 慢慢地轻轻取出套管针,确保肿瘤在第四个乳腺脂肪垫内保持深度。
    1. 皮下移植:
      1. 打开阀门到鼻锥,将鼠标的鼻子放在鼻锥中,鼠标的腹侧面向下朝向吸收垫。
      2. 刮下右腿,背部的背部从后肢到前肢
      3. 用betadine擦拭剃光皮肤。
      4. 用无菌酒精制剂垫擦去betadine。
      5. 用镊子将背部的背部向上拉,用剪刀(4-5毫米)水平切割小切口。
      6. 使用无菌剪刀或镊子在皮肤下方形成一个小口袋,并将肿瘤碎片放在皮肤下面。确保将肿瘤片断距开口至少3mm。
    2. 用镊子抓住初始手术口的两端,拉起并调整手和镊子,以便用一个镊子抓住切口的边缘。同时仍然拉起皮肤,缝合切口关闭。避免装订到鼠标的体腔。
    3. 将鼠标转移到37°C保温箱上的临时,干净的笼子上。密切监视动物,直到恢复意识。
    4. 继续对剩下的四只小鼠的肿瘤植入物,手术后转移到加温笼。当所有小鼠恢复意识并正常移动时,可以将其转移到标准的笼子上。
    5. 当第五只小鼠完成手术时,您可以将另外5只小鼠转移到异氟醚室麻醉。
    6. 完成后,关闭异氟烷机,用70%乙醇喷洒所有表面,然后擦拭,然后进行防腐处理,如Clidox。
    7. 通过在水中冲洗清洁所有仪器,然后用手术器械清洁剂清洗。高压釜消毒。
    8. 7-14天后取出订书钉。

  4. 植入雌激素颗粒
    注意:请查看视频,了解有关程序的其他信息(视频4)。

    <! - flashid2100v150开始 - >
    视频4.皮下移植肿瘤移植
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    如果使用乳腺癌细胞或患有雌激素依赖性的患者来源的乳腺癌肿瘤,则需要用雌激素补充肿瘤生长。推荐使用持续释放超过60-90天的雌激素颗粒。
    1. 该过程需要麻醉,并且可以在移植肿瘤或细胞接种之前或之后进行。
    2. 根据您的机构IACUC协议麻醉与异氟烷的鼠标。
    3. 将头发从肩胛骨上方的颈部背侧剃掉。使用脱毛膏不推荐用具有SCID背景的免疫缺陷型模型。
    4. 用betadine棉签擦拭暴露的皮肤,并用无菌酒精垫擦拭。
    5. 做一个直径相等于切片的切口。
    6. 使用镊子拉开切口一侧的皮肤,并用一对镊子或剪刀将水平方式制成口袋,距离切口部位约1厘米。
    7. 使用镊子将雌激素颗粒插入口袋。
    8. 缝合或胶合切口(一些老鼠不会忍受他们的颈部的钉子,并将其刮掉,导致雌激素颗粒的损失)。
    9. 7-14天后取出订书钉。

  5. 肿瘤组织的低温保存
    1. 准备冷冻介质:培养基+ 10%FBS +无菌10%DMSO。
    2. 将无菌收获的肿瘤组织置于无菌培养皿中,并用约5ml的含有1%青霉素 - 链霉素的无菌PBS冲洗。
    3. 可见地检查组织以从肿瘤标本中小心地去除和丢弃任何坏死和非肿瘤组织
    4. 使用镊子和手术刀,将肿瘤组织切割成3×3×3 mm片段;取决于肿瘤类型,一些组织可能通过用镊子分开分开而不是用手术刀切割更容易解剖。
    5. 标记具有肿瘤组织ID的低温小瓶,收获日期,加入1ml冷冻培养基
    6. 将单个肿瘤片段放入低温小瓶中,拧紧帽子并置于慢速冷冻设备中
    7. 将盖子放在冷冻设备上并置于-80°C冷冻室过夜。肿瘤组织以0.5°C/min的速度冷冻至-20°C,1°C/min至-80°C。
    8. 过夜冷冻后,将低温小瓶储存在液氮冷冻箱中,并记录冻存温度的位置和日期
  6. 肿瘤测量和药物治疗(口服管饲和腹膜内注射)
    1. 应使用卡尺(长度,宽度和高度)每周测量2〜3次肿瘤生长。
    2. 以允许进入肿瘤的方式轻轻地限制鼠标。
    3. 将卡尺带到肿瘤的最长侧以测量长度。轻轻地将卡钳挤压在一起,直到肿瘤的边界在卡钳钳口内。
    4. 将卡尺放在肿瘤较窄的一侧以测量宽度。
    5. 高度测量可以通过尝试使用卡尺来测量肿瘤的高度。或者,塑料尺可以放置在与肿瘤底部平齐并且肿瘤的高度读取。
    6. 适当记录长度,宽度和高度。
      注意:对于大多数一致的结果,同一个人应在研究期间测量肿瘤。否则,可能会观察到变异性,因为每个人在测量期间可能会不同地挤压肿瘤。
    7. 可以通过椭圆体积公式(π/6×L×W×H)或体积公式(L x W x H)(Tomayko和Reynolds,1989)来计算肿瘤体积。
    8. 开始药物治疗的肿瘤体积取决于实验的目标,但通常肿瘤应至少为100毫米以上,以确保肿瘤生长。如果实验的目标是测试"预防"肿瘤生长,则应调整药物治疗时间表。
      1. 对于在已建立的肿瘤进行的实验,使用近似肿瘤体积平均值为150mm 3的药物治疗起始大小。当一群动物具有150mm 3的平均肿瘤大小时,将小鼠随机分成治疗组。肿瘤小于115 mm 3的动物不包括在第一个队列中,并且将被随机分配到第二组小鼠中,并在以后的研究中报名参加。
        注意:肿瘤生长的速率和均匀性完全取决于肿瘤细胞系或肿瘤移植物的性质。在一些实验中,所有移植的小鼠将能够同时开始治疗,并且在一些实验中,可能需要将动物分成几个在不同日期进行研究的群组。
      2. 随机化后,根据其分配的治疗组,用载体或药物治疗小鼠
      3. 剂量,给药方案(每周,每天,每天两次,等等)和载体由化合物确定。这些信息通常在已出版的文献或提供该化合物的制药公司提供。
    9. 口服管饲。
      1. 称重动物以确定所需的平均体重和剂量/体积。可以在小鼠中灌胃的最大体积为200μl。  
      2. 通过连接到1cc注射器来准备22号管饲枪。
      3. 通过用拇指和食指抓住肩膀上的皮肤使得前腿伸出到侧面,保持前脚不要将管饲管推开,将动物限制在直立的位置。
      4. 通过轻轻地将头部向后延伸(头部的这个延伸部分通过颈部和食道创建一条直线)将动物的头部保持在位。
      5. 管饲针用于测量从嘴到小鼠的最后一个肋(胃)的距离。这是几乎要插入管饲针以确保有效的程序。
      6. 将针的灯泡插入口角。将灯泡重新放置在中心,沿着嘴巴的顶部运行。
      7. 使用针作为杠杆,轻轻地向后(直立)鼠标的头部和鼻子。 (限制手保持直立,不弯曲。) 
      8. 慢慢地将管饲针沿着食道喂给预先测量的距离。如果感觉到阻力,请勿进一步推针。如有必要,请将针头缩回,直到灯泡在嘴部前方(但不是外面)并重新插入。不要尝试插入3次以上。如果3次尝试后插入失败,必须允许动物休息2小时。在休息期之后,可以再次尝试插入3次。如果第二轮不合格,则在继续口头管治之前,应咨询参加兽医的事宜
      9. 推进针头,使头部和鼻子进一步向后或直立。
      10. 在推动柱塞之前,请确保针头完全插入到预先测量的目标点。
      11. 推动柱塞将内容物释放到动物的胃中。
      注意:在研究中可以在多只动物上使用管饲针。但是,治疗之间应该改变针头。
    10. 腹膜内(IP)注射
      1. 为了确定针头的入口点在膝盖上方的腹部穿过假想线。针将沿着动物右侧的这条线插入并靠近中线。在女性动物中,进入点是最后一个乳头的颅内和稍微内侧。将针插入鼠标右侧可避免盲肠。
      2. 要限制鼠标:用拇指和食指抓住肩膀上的皮肤。倾斜头部,使其朝下,腹部暴露。要执行IP注入,鼠标必须有良好的约束,以便在此过程中无法移动。一旦针进入腹部,就避免了器官的创伤
      3. 将27号针头以大约30度的角度插入腹部。针的轴应进入约半厘米的深度。
      4. 在注射前,抽出来确保针头没有穿透血管,肠或膀胱。
        1. 绿棕色抽出物表示针刺入肠。
        2. 黄色吸出物表示针刺入膀胱。
        3. 如果吸入任何液体,您的溶液将被污染,必须将其丢弃,并用新的注射器和针头重复操作。
        4. 如果没有吸入液体,可以注射。
      5. 拔出针头并将鼠标移回笼子。

数据分析

对于药物治疗实验,基于肿瘤大小将小鼠随机分为对照组或治疗组。每个实验/治疗组所需的小鼠数量通过功率分析来确定。在治疗前和治疗后纵向监测肿瘤体积。生化统计学家对治疗状态不了解,以缓解对统计分析引入的偏倚。线性混合效应模型用于测试跨治疗组的药物反应的显着差异,如果存在不良生长的异种移植/PDX受试者,则利用XenoCat建模框架来增加统计学功能(Laajala等人, 2012)。引用适当的模型对比来正式测试药物反应中的协同作用或拮抗作用。 Bonferroni校正被应用于控制家庭错误率。

笔记

  1. 在用细胞系运行大型异种移植研究之前,建议进行一项小型试验性研究以确定肿瘤的采集率。如果摄取率小于100%,这将有助于确定需要的额外小鼠的数量。
  2. 细胞系异种移植物的成功和可行性可能受到收获时融合的影响。如上所述,当细胞处于指数生长阶段时,最好收获。如果细胞系异种移植物的肿瘤活力差,也可以将Matrigel加入到细胞培养基注射混合物中,以提供植入物的富集基质环境。
  3. 使用患者来源的异种移植模型,植入物高度依赖于肿瘤类型。转移性乳腺癌可以获得25%的收获率,但是在我们的研究中,治疗原始乳腺癌的接种率接近5%。

致谢

我们要感谢Bryn Eagleson和VARI Vivarium员工的专业知识。这里描述的方案用于由乳腺癌研究基金会,Muskegon诱惑表和Van Andel基金会支持的多项研究。

参考文献

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  2. DeRose,YS,Gligorich,KM,Wang,G.,Georgelas,A.,Bowman,P.,Courdy,SJ,Welm,AL和Welm,BE(2013)。  人乳腺癌的患者衍生模型:体外和在肿瘤生物学和翻译医学中的体内应用 Curr Protoc Pharmacol 第14章:Unit14 23.
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引用:Tovar, E. A., Essenburg, C. J. and Graveel, C. (2017). In vivo Efficacy Studies in Cell Line and Patient-derived Xenograft Mouse Models. Bio-protocol 7(1): e2100. DOI: 10.21769/BioProtoc.2100.
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