Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2013 Apr 1.
Published in final edited form as: Nat Protoc. 2012 Mar 22;7(4):749–755. doi: 10.1038/nprot.2012.030

An isolated tumor perfusion model in mice

Annique M M J Duyverman 1,2, Mitsutomo Kohno 1,2, Sylvie Roberge 1, Dai Fukumura 1, Dan G Duda 1, Rakesh K Jain 1
PMCID: PMC3375334  NIHMSID: NIHMS379980  PMID: 22441293

Abstract

The role of stromal cells in the tumor microenvironment has been extensively characterized. We and others have shown that stromal cells may participate in several steps of the metastatic cascade. This protocol describes an isolated tumor perfusion model that enables studies of cancer and stromal cell shedding. It could also be used to study the effects of therapies interfering with the shedding of tumor cells or fragments, circulating (stem) cells or biomarkers. Primary tumors are grown in a microenvironment in which stromal cells express GFP ubiquitously. Tumors are implanted orthotopically or can be implanted ectopically. As a result, all tumor-associated stromal cells express GFP. This technique can be used to detect and study the role of stromal cells in tumor fragments within the circulation in mice. Studying the role of stromal cells in circulating tumor fragments using this model may take 2–10 weeks, depending on the growth rate of the primary tumor.

INTRODUCTION

Neoplastic tumor cells and host-derived stromal cells coexist in carcinomas. Clinical and preclinical evidence supports the contribution of these nonmalignant stromal cells to the development of tumors1-6. Many studies have shown that solid tumors actively recruit these stromal cells to create a microenvironment that promotes primary tumor growth and dissemination7-10. Moreover, this process is likely to be repeated when tumors colonize secondary sites during metastatic growth.

A tumor cell from the primary tumor can lead to established metastases when it successfully completes each of the sequential and interrelated steps of the metastatic cascade. Each of these steps can be rate limiting, and failure to complete a step can abrogate the entire process11. Recent studies have shown that the host-derived stroma may increase the efficiency of this complex system. Among the stromal cell types that have been linked to tumor promotion and progression are endothelial cells, pericytes, fibroblasts and various bone marrow–derived cells, including splenocytes, macrophages, neutrophils, mast cells, myeloid-derived suppressor cells and mesenchymal stem cells6,12,13. Metastatic cells could reside in the lungs awaiting oncogenic activation14, or could home to pre-existing niches created by inflammation and immune cell or fibroblast accumulation15-18. In addition, it has been shown that metastatic cells proliferate intravascularly before extravasation into the lung tissue19.

Previous applications of the protocol

Studies reported more than 30 years ago showed that cancer cell clumping in circulation increases metastasis20,21. These clumps may be emboli, formed in circulation because of interactions with immune cells11,22,23. Indeed, the injection of emboli-containing tumor cells increases the efficiency of metastasis. However, the clumps may also be fragments consisting of tumor cells that carry ‘passenger’ stromal cells from the primary site. By using the techniques described in this protocol in combination with two spontaneous metastases models—one using skin transplantation by transient parabiosis and a second using carcinoma-associated fibroblasts to selectively deplete stromal cells24,25—we have recently shown that these host-derived cells increase tumor cell viability in circulation and serve as a provisional stroma in the secondary site and increase the cancer’s metastatic efficiency26.

Overview of the technique

The technique described in this protocol can be used to study circulating tumor cells originating from any primary tumor. Tumors can be implanted orthotopically or ectopically, and non-tumor-bearing mice are used as a control. Here we use the renal perfusion model in combination with primary tumor implantation in the renal capsule. Implanting tumors at different (orthotopic) sites might provide additional insights in assays of mestastasis, and they could be adapted from previously established isolated tumor models. These include transplanted and spontaneous breast cancer, ovarian cancer and liver cancer models27-30. The experimental design of the protocol used is depicted in Figure 1. To study tumor cells and fragments shed by the primary tumor, we implant fluorescent-labeled tumor cells in the kidney capsule. After 12 d, we use the isolated kidney-tumor perfusion model to collect tumor cells and fragments that were shed by the primary tumor. By monitoring blood pressure in the carotid artery and adjusting the perfusion rate through the jugular vein, we are able to collect tumor perfusate for up to an hour, yielding 3–4 ml of blood per mouse.

Figure 1.

Figure 1

Open in a new tab

Design of the experiment. LLC1-dsRed tumor cells are implanted in the renal capsule. When the tumor size is 8 mm, the renal vein is cannulated, and tumor cells and fragments are collected by filtration of the perfusate using a peristaltic pump and artificial blood perfusion. Reproduced with permission from ref. 26. All animal procedures were performed according to the guidelines of public health service policy on humane care of laboratory animals and in accordance with an approved protocol by the Institutional Animal Care and Use Committee of MGH.

Comparison with other techniques

Several in vitro assays have been developed to study host–tumor cell interactions related to progression and invasion31,32, including the chemoinvasion assay using the reconstituted BD Matrigel in Boyden blind-well chambers. It can be applied to detect the migratory activity associated with matrix degradation and can also be adapted to study the selective degrading activity on different matrix substrates. The advantage of this in vitro assay is the ease of controlling different parameters in the experiment. At the same time, the lack of in vivo microenvironmental factors may confound the results. Many in vivo models have been used to examine the role of stromal components within the primary tumor microenvironment. Human mammary tissue can be reconstructed in a mouse model to study cross talk between tissue stroma and the epithelium, as well as factors involved in breast stem cell biology of tumor initiation and progression33. However, there is a paucity of techniques for studying and visualizing primary tumor stromal cells in different steps of the metastatic process. We have developed an experimental protocol that can be used to quantitatively study the contribution of stromal cells to tumor fragment survival within the circulation. We describe the isolated tumor perfusion model to study the viability of tumor cells in circulating tumor fragments. This model was originally developed by Gullino, and later adapted by us for various molecular and cellular studies of cancer28-30,34,35. We have used this model to unravel the role of ‘passenger’ fibroblasts in tumor metastasis to the lungs26. However, other tumor stromal cells could potentially be studied using this protocol.

Advantages and limitations

The key advantage of our protocol over other existing methods is that one can gain the ability to image and quantify in vivo all cells and fragments that are shed by the kidney, liver, ovary and inguinal gland during metastasis. Alternatively, one could use the collected cells/fragments for consecutive in vivo experiments in which the effect of drugs on tumor cell/clump shedding could be studied.

The major limitation of this technique is that animals have to be exsanguinated. In addition, a large number of mice are required to study the effects of therapy over time.

The renal perfusion model shown here is an ectopic tumor model, where tumors are implanted surgically in the renal capsule. However, other organs or tissues could, in principle, be used to collect efferent cells.

A further potential disadvantage is that the studies require GFP+ or other transgenic immunocompetent mice, which can only be used with certain (syngeneic) tumor lines.

MATERIALS

Inline graphic CRITICAL All reagents and equipment can be substituted with appropriate alternatives from other manufacturers.

REAGENTS

  • PBS (Cellgro, cat. no. 20-031-CV)

  • Buprenorphine hydrochloride (0.3 mg ml−1; MGH pharmacy, cat. no. 716510) ! CAUTION Buprenorphine hydrochloride is a poison, and may cause prolonged respiratory depression. Wear protective clothing to avoid contact or inhalation. Buprenorphine is a controlled substance and should be handled according to relevant rules of the host institutions.

  • Ethanol (70% (vol/vol); Pharmco, cat. no. 111000190)

  • Fluorescent-labeled LLC1 cells (e.g., dsRed-labeled cells; ATTC, cat. no. CRL-1642)

  • Hank’s balanced salt solution (1×; Gibco, cat. no. 14170)

  • Heparin sodium (1,000 USP Units ml−1; APP Pharmaceuticals, cat. no. 504011)

  • Ketamine (100 mg ml−1; MGH pharmacy) and xylazine (10 mg ml−1; Webster, cat. no. 200204.00) mixture per kg of body weight)

  • Mice, male Actb-GFP/C75BL/6, 6–10 weeks of age ! CAUTION All animal studies must be reviewed and approved by the institutional animal care and use committees to ensure that they conform to relevant ethics regulations.

  • Cells for transplantation, e.g., LLC1-dsRed tumor cells

  • Cyanoacrylate glue (Krazy Glue) ! CAUTION Avoid contact with skin, and wear gloves when handling.

  • Formaldehyde

  • Paraformaldehyde

  • Sodium chloride

  • dH2O

EQUIPMENT

  • 5-0 Ethibond sutures (Ethicon, cat. no. X698G)

  • Polyethylene 10 + 50 tubing (PE10, PE50; Becton Dickinson, cat. nos. 427401 and 427411)

  • Applicator tips (Owens and Minor, cat. no. 5937-W0D1002)

  • Bright-field microscope

  • Caliper (Roboz Surgical Instrument, cat. no. RS-6466)

  • Clipper (Webster, cat. no. 78997-010)

  • Fluorescence confocal or multiphoton microscope (e.g., Olympus). For confocal microscopy, use the Argon laser, 488 nm excitation and an emission filter of 505–525 nm to detect GFP signal, and the HeNe laser, 543 nm excitation and an emission filter of 565–595 nm to detect dsRed signal. For two-photon microscopy, use 840 nm excitation and an emission filter of 515–555 nm (for GFP) or/and an emission filter of 575–645 nm (for dsRed)

  • Forceps (Roboz Surgical Instrument, cat. no. RS-5153, RS 5150, RS-5132)

  • Gas exchanger (Becton Dickson)

  • Heating pad (Shore Line, cat. no. 712.0000.04)

  • Hemostatic forceps (9-inch, jaw length 6 cm; Roboz Surgical Instrument, cat. no. RS-7679)

  • Micro aneurysm clip, straight, plus clip applier (Roboz Surgical Instrument, cat. no. RS-5420 + RS-8140)

  • Micro forceps (2×; F.S.T. cat. no. 11063-07)

  • Micro forceps (2×; Roboz Surgical Instrument, cat. no. RS-5069)

  • Micro scissors (Tiemann, cat. no. 160-147)

  • Needle holder (14 cm; Roboz Surgical Instrument, cat. no. RS-6412)

  • Peristaltic pump (Ismatec cat. no. 7618-3)

  • Permanent marker (Staedtler, cat. no. 342-9)

  • Pressure transducer (Gould INC)

  • Reservoir of Oxyglobin (Biopure, cat. no. HBOC-301)

  • Scissors (Roboz Surgical Instrument, cat. no. RS-5840, RS-5883)

  • Silastic tubing (Fisher, cat. no. 11-18915C)

  • Three-way stopcock (Cole-Parmer, cat. no. 30600-02)

  • Surgical blade (no.10; Fisher Scientific, cat. no. 08-916-5A)

  • Scalpel handle (Roboz Surgical Instrument, cat. no. RS-9843)

  • Syringe (1 cc with a 26-G needle for anesthesia; Fisher Scientific, cat. no. 14-823-2E)

  • 30-G needle for injecting tumor cells

  • 6-0 silk sutures

REAGENT SETUP

Paraformaldehyde, 4% (wt/vol) Add 180 ml of 1× PBS to 120 ml of 10% (wt/vol) formaldehyde. Inline graphic CRITICAL This solution has a short shelf life (< 1 week); therefore, it should be freshly prepared on the day of the experiment.

Buprenorphine hydrochloride Dissolve 1 ml of the 0.3 mg ml−1 stock solution in 30 ml of 0.9% (wt/vol) sodium chloride. Store at 20 °C for up to 3 months.

PBS (1×) Add 100 ml of 10× PBS to 900 ml of dH2O. Store at 20 °C for up to 9 months.

Ethanol, 70% (vol/vol) Mix 1.7 liters of dH2O and 1 gallon of 100% ethanol. Store at 20 °C in a closed container.

Heparin Dilute 0.3 ml of stock solution in 30 ml of sodium chloride (freshly prepare for each experiment).

PROCEDURE

Tumor implantation in the left kidney

1| Anesthetize a male Actb-GFP/C57BL/6 mouse, 6–10 weeks of age, using an intramuscular injection of 0.4 ml of ketamine/xylazine and place it on a heating pad.

2| Position the mouse on the heating pad with the left flank facing the surgeon. Make an 8-mm incision through the skin and muscle layers approximately 1 cm lateral to the spine and 3 mm dorsal to the lowest ribs. Externalize the left kidney using applicator tips.

? TROUBLESHOOTING

3| Create a small pocket under the renal capsule using the tip of the micro syringe. Inject 1 × 106 LLC1-dsRed tumor cells in 0.1 ml of Hank’s balanced salt solution in the renal capsule using a 30-G needle.

? TROUBLESHOOTING

4| Remove any liquids that are leaking out of the capsule using an applicator tip.

5| Close the muscle layers and skin using 5-0 Ethibond sutures. Keep the mice in appropriate cages under sterile conditions and allow the tumors to grow to 8 mm.

6| At the preferred time point (e.g., 2 weeks after implantation of LLC1 tumors), anesthetize the mouse using an intramuscular injection of 0.4 ml of ketamine/xylazine and keep it on a heating pad.

7| Open the abdomen by laparatomy to inspect the location of the kidney tumor. Reposition the organs and cover the abdomen with skin and a sponge before moving to the next step.

? TROUBLESHOOTING

cannulation of the carotid artery

8| Position the mouse on its back, with the mouse’s head facing the surgeon. Fix the head by taping it across the nose and fix the body by taping across the paws and the sternum (Fig. 2a).

Figure 2.

Figure 2

Open in a new tab

Cannulation of the carotid artery. (a) Position the mouse on its back, head facing the surgeon. Fix the head by taping it across the nose and fix the body by taping across the paws and the sternum. (b) Make a midline incision under the chin, exposing the salivary glands. Expose the trachea by separating the salivary glands sideways using the micro forceps. (c) Apply the hemostatic clamp onto the carotid artery caudal to the loose knot. All animal procedures were performed according to the guidelines of public health service policy on humane care of laboratory animals and in accordance with an approved protocol by the Institutional Animal Care and Use Committee of MGH.

9| Make a midline incision under the chin, exposing the salivary glands. Expose the trachea by separating the salivary glands sideways using the micro forceps (Fig. 2b).

10| Dissect the muscles overlying the carotid artery using microdissection forceps and dissection microscope.

Inline graphic CRITICAL STEP It is important to remove all the connective tissue surrounding the carotid artery, as this facilitates cannulation.

11| Loop two 6-0 silk sutures around the artery, 5 mm apart, and then close the cranial ligature. Make a loose knot in the caudal suture (Fig. 2c).

12| Apply the hemostatic clamp onto the carotid artery caudal to the loose knot (Fig. 2c).

13| Connect the heparin-filled PE10 tubing to a three-way stopcock and pressure transducer through a 30-G needle. While holding the cranial suture, cut across one-third of the carotid vessel wall with microscissors, and then insert the heparin-filled PE10 tubing into the vessel toward the hemostatic clamp.

Inline graphic CRITICAL STEP Do not section the vessel completely, as this makes inserting the tubing impossible. Release the clamp and insert the tubing 3 mm beyond the clamp.

? TROUBLESHOOTING

14| Close the suture around the vessel and tubing to secure the tubing. Total tubing insertion is 8 mm. At the other end of the PE tubing, insert a 30-G needle connected to a three-way stopcock and the pressure transducer. Add a drop of Krazy Glue to fix the tubing and vessel.

? TROUBLESHOOTING

cannulation of the jugular vein

15| Move to the right side of the trachea to cannulate the jugular vein.

16| Remove the connective tissue surrounding the jugular vein for 8 mm. If any vessel branches off the vein, then ligate it with two silk sutures and cut in between.

Inline graphic CRITICAL STEP Dissect the jugular vein in its anatomical position to avoid rupture of this delicate vein; it has a much thinner wall than the carotid artery.

17| Loop two 6-0 silk sutures around the vein, 5 mm apart, and then close the cranial ligature. Make a loose knot in the caudal suture.

18| Apply the hemostatic clamp onto the jugular vein caudal to the loose knot.

19| Connect the PE50 tubing to a reservoir of Oxyglobin. While holding the cranial suture, cut across one-third of the vessel wall using microscissors and insert the PE50 tubing in the direction of the hemostatic clamp. Release the clamp and insert the tubing 3 mm beyond the clamp. Close the suture around the vessel and tubing, and add a drop of Krazy Glue to secure the tubing. The total tubing insertion length is 8 mm.

Inline graphic CRITICAL STEP As there is not yet any marked loss of circulating volume, perfuse at very low rates.

? TROUBLESHOOTING

Kidney perfusion

20| Reposition the mouse on its back with the tail facing the surgeon.

Inline graphic CRITICAL STEP Do not disturb the tubing inserted in the carotid artery or jugular vein while repositioning the animal.

21| Access the abdominal cavity through the previously performed laparatomy.

22| Move and hold all the organs to the right side using a wet sponge to clear the field for the cannulation of the left renal vein.

23| Locate the left adrenal vein and testicular vein draining into the renal vein and ligate both to avoid rupture. Use two 6-0 silk sutures to ligate the branches and cut in between to create a space to move the renal vein around.

24| Dissect the connective tissue around the renal vein to locate the renal artery, which is behind the vein, and then separate the two.

25| Loop two 6-0 silk sutures around the renal vein. Close the proximal one and keep a loose knot in the distal one for tube insertion. While holding the closed suture, cut 30% of the vein across and insert a 5-cm-long, heparinized PE10 as close to the kidney as possible. Close the suture to fix the tubing and add a drop of Krazy Glue at the point of insertion.

? TROUBLESHOOTING

26| Collect the dripping perfusate into the 15-ml collection tube kept on ice, and increase the Oxyglobin flow to maintain a physiological pressure between 60 and 100 mm Hg. Oxygenation of the perfusate solution is achieved by a gas exchanger in which the perfusate is led through 16 feet of silastic tubing while being equilibrated with warm humidified 95% O2 and 5% CO2. During the experiment, keep the perfusate in a heated bath and lead it through the gas exchanger and then into the jugular vein into the mouse’s circulation.

? TROUBLESHOOTING

27| Cover the open abdomen with a wet sponge for the remaining time of the experiment, up to 1 hour.

? TROUBLESHOOTING

? TROUBLESHOOTING

Troubleshooting advice can be found in Table 1.

TABLE 1.

Troubleshooting table

Step Problem Possible reason Solution
2 Inability to exteriorize the
kidney		 Incision is not in the correct
place		 Reposition/enlarge the incision
3 Injected cells leak out Pocket is not large enough Exclude the mouse from procedure and create larger
pockets without rupturing the renal capsule
7 Tumor is growing close to the
kidney vein or invades more
than 30% of the kidney		 Blood flow from the kidney into
the renal vein is impaired by the
tumor		 Exclude the mouse from the procedure
13 Difficulty inserting the tubing
into the vessel		 Tubing is not shaped correctly Cut the tubing diagonally for easier insertion
14 Cannula leak Knot not secured tightly Leave the inferior knot in place, loop and knot an
additional suture proximally and tighten securely
Obstructed tubing Blood clot in the tubing Heparinize the tubing with 0.05 M heparin
14, 19, 25 Extravasation of the tubing Tubing is disconnected during
repositioning of the animal		 Securely tighten the knots in sutures and apply glue
26 Collected cells are not viable Cell death Keep the perfusate on ice during the experiment
27 Premature death of the mouse Hyper/hypotension Carefully monitor blood pressure during the
collection of perfusate and adjust Oxyglobin flow
Open in a new tab

Inline graphic TIMING

Steps 1 and 2, anesthesia and positioning of the mouse: 10 min

Steps 3 and 4, tumor cell injection: 3 min

Step 5, muscle and skin closure: 10 min

Steps 6–14, cannulation of the carotid artery: 10 min

Steps 15–19, cannulation of the jugular vein: 20 min

Steps 20–26, cannulation of renal artery: 20 min

Step 27, collection of perfusate: 1 h

ANTICIPATED RESULTS

Figure 2 shows some typical results of the renal perfusion model. Heterogeneous tumor and stromal cell clumps can be detected in the filtered perfusate using a fluorescence microscope (Fig. 2). These harvested clumps and single cells can be further studied for viability, composition or expression of specific proteins of interest. In our study, the vast majority (~81%) of the shed dsRed+ cancer cells were single cells (Fig. 3a). However, we also collected small tumor clumps (≤200 μm in diameter), and all tumor clumps composed of six or more cells contained GFP-expressing host cells (Fig. 3a). In addition, caspase 3 and caspase 7 activation—a measure of apoptosis—was detectable in most (~88%) of the single or doublets of cancer cells at the time of shedding. In contrast, the heterotypic cell clumps contained almost twice as many viable cancer cells (22.8 ± 4.5%, P < 0.05; see Fig. 3b). The clumps can be used for consecutive in vitro or in vivo experiments to study the effect of antimetastatic drugs on tumor cell clumping and viability (Fig. 4).

Figure 3.

Figure 3

Open in a new tab

Representative images of tumor cell clumps retained on a 40-mm mesh. (a,b) Clumps filtered from tumor perfusate. Red, LLC1-dsRed cells; green, GFP+ host-derived cells from the Actb-GFP/C57BL/6 mice.

Figure 4.

Figure 4

Open in a new tab

Size and viability of circulating metastatic cancer cells. (a) Histogram of the composition of shed tumor cells/clumps obtained from the renal perfusion experiment (n = 5 mice). The majority of shed cancer cells were single or doublets. Host-derived GFP+ cells were present in all large clumps consisting of more than 4–5 cells (*P < 0.05). (b) Representative fluorescence multiphoton laser scanning microscopy image of a heterogeneous clump, shed by a tumor, using the isolated renal perfusion model (green, GFP+ stromal cells; red, dsRed+ tumor cells). Image is 420 μm across. Reproduced with permission from ref. 26. All animal procedures were performed according to the guidelines of public health service policy on humane care of laboratory animals and in accordance with an approved protocol by the Institutional Animal Care and Use Committee of MGH.

ACKNOWLEDGMENTS

The work of the authors is supported by US National Cancer Institute grants P01-CA80124, R01-CA115767, R01-CA85140, R01-CA126642 and T32-CA73479 (R.K.J.), R01-CA96915 (D.F.), R21-CA139168 and R01-CA159258 (D.G.D.) and Federal Share Proton Beam Program grants (R.K.J., D.F. and D.G.D.); Department of Defense Innovator Award W81XWH-10-1-0016 (R.K.J.) and Predoctoral Fellowship W81XWH-06-1-0781 (A.M.M.J.D.); American Cancer Society grant RSG-11-073-01TBG (D.G.D.); and Stichting Michael Van Vloten Fonds and the Stichting Jo Kolk (A.M.M.J.D.). We acknowledge the outstanding technical assistance of J. Kahn and P. Huang with animal models.

Footnotes

AUTHOR CONTRIBUTIONS D.G.D., D.F. and R.K.J. designed the studies; A.M.M.J.D., S.R. and M.K. performed the experiments; D.G.D., D.F., A.M.M.J.D., M.K. and R.K.J. analyzed the data; and A.M.M.J.D., D.G.D., D.F. and R.K.J. edited the manuscript.

COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests.

References

  • 1.Barcellos-Hoff MH, Ravani SA. Irradiated mammary gland stroma promotes the expression of tumorigenic potential by unirradiated epithelial cells. Cancer Res. 2000;60:1254–1260. [PubMed] [Google Scholar]
  • 2.Bhowmick NA, et al. TGF-β signaling in fibroblasts modulates the oncogenic potential of adjacent epithelia. Science. 2004;303:848–851. doi: 10.1126/science.1090922. [DOI] [PubMed] [Google Scholar]
  • 3.Coussens LM, Werb Z. Inflammation and cancer. Nature. 2002;420:860–867. doi: 10.1038/nature01322. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4.Jacobs TW, Byrne C, Colditz G, Connolly JL, Schnitt SJ. Radial scars in benign breast-biopsy specimens and the risk of breast cancer. N. Engl. J. Med. 1999;340:430–436. doi: 10.1056/NEJM199902113400604. [DOI] [PubMed] [Google Scholar]
  • 5.Elenbaas B, et al. Human breast cancer cells generated by oncogenic transformation of primary mammary epithelial cells. Genes Dev. 2001;15:50–65. doi: 10.1101/gad.828901. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6.Bouvet M, et al. In vivo color-coded imaging of the interaction of colon cancer cells and splenocytes in the formation of liver metastases. Cancer Res. 2006;66:11293–11297. doi: 10.1158/0008-5472.CAN-06-2662. [DOI] [PubMed] [Google Scholar]
  • 7.Bhowmick NA, Neilson EG, Moses HL. Stromal fibroblasts in cancer initiation and progression. Nature. 2004;432:332–337. doi: 10.1038/nature03096. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 8.Olumi AF, et al. Carcinoma-associated fibroblasts direct tumor progression of initiated human prostatic epithelium. Cancer Res. 1999;59:5002–5011. doi: 10.1186/bcr138. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 9.Orimo A, et al. Stromal fibroblasts present in invasive human breast carcinomas promote tumor growth and angiogenesis through elevated SDF-1/CXCL12 secretion. Cell. 2005;121:335–348. doi: 10.1016/j.cell.2005.02.034. [DOI] [PubMed] [Google Scholar]
  • 10.Tlsty TD. Stromal cells can contribute oncogenic signals. Semin. Cancer Biol. 2001;11:97–104. doi: 10.1006/scbi.2000.0361. [DOI] [PubMed] [Google Scholar]
  • 11.Fidler IJ. The pathogenesis of cancer metastasis: the ‘seed and soil’ hypothesis revisited. Nat. Rev. Cancer. 2003;3:453–458. doi: 10.1038/nrc1098. [DOI] [PubMed] [Google Scholar]
  • 12.Fukumura D, et al. Tumor induction of VEGF promoter activity in stromal cells. Cell. 1998;94:715–725. doi: 10.1016/s0092-8674(00)81731-6. [DOI] [PubMed] [Google Scholar]
  • 13.Joyce JA, Pollard JW. Microenvironmental regulation of metastasis. Nat. Rev. Cancer. 2009;9:239–252. doi: 10.1038/nrc2618. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14.Podsypanina K, et al. Seeding and propagation of untransformed mouse mammary cells in the lung. Science. 2008;321:1841–1844. doi: 10.1126/science.1161621. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15.Hiratsuka S, et al. MMP9 induction by vascular endothelial growth factor receptor-1 is involved in lung-specific metastasis. Cancer Cell. 2002;2:289–300. doi: 10.1016/s1535-6108(02)00153-8. [DOI] [PubMed] [Google Scholar]
  • 16.Kaplan RN, et al. VEGFR1-positive haematopoietic bone marrow progenitors initiate the pre-metastatic niche. Nature. 2005;438:820–827. doi: 10.1038/nature04186. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 17.Kim S, et al. Carcinoma-produced factors activate myeloid cells through TLR2 to stimulate metastasis. Nature. 2009;457:102–106. doi: 10.1038/nature07623. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 18.van Deventer HW, et al. C-C chemokine receptor 5 on pulmonary fibrocytes facilitates migration and promotes metastasis via matrix metalloproteinase 9. Am. J. Pathol. 2008;173:253–264. doi: 10.2353/ajpath.2008.070732. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 19.Al-Mehdi AB, et al. Intravascular origin of metastasis from the proliferation of endothelium-attached tumor cells: a new model for metastasis. Nat. Med. 2000;6:100–102. doi: 10.1038/71429. [DOI] [PubMed] [Google Scholar]
  • 20.Liotta LA, Saidel MG, Kleinerman J. The significance of hematogenous tumor cell clumps in the metastatic process. Cancer Res. 1976;36:889–894. [PubMed] [Google Scholar]
  • 21.Fidler IJ. The relationship of embolic homogeneity, number, size and viability to the incidence of experimental metastasis. Eur. J. Cancer. 1973;9:223–227. doi: 10.1016/s0014-2964(73)80022-2. [DOI] [PubMed] [Google Scholar]
  • 22.Ruiter DJ, van Krieken JH, van Muijen GN, de Waal RM. Tumour metastasis: is tissue an issue? Lancet Oncol. 2001;2:109–112. doi: 10.1016/S1470-2045(00)00229-1. [DOI] [PubMed] [Google Scholar]
  • 23.Sahai E. Illuminating the metastatic process. Nat. Rev. Cancer. 2007;7:737–749. doi: 10.1038/nrc2229. [DOI] [PubMed] [Google Scholar]
  • 24.Duyverman AM, Steller EJ, Fukumura D, Jain RK, Duda DG. Studying carcinoma-associated fibroblast involvement in cancer metastasis in mice. Nat. Protoc. 2012;7:756–762. doi: 10.1038/nprot.2012.031. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 25.Duyverman AMMJ, Kohno M, Duda DG, Jain RK, Fukumura D. A transient parabiosis skin transplantation model in mice. Nat. Protoc. 2012;7:763–770. doi: 10.1038/nprot.2012.032. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 26.Duda DG, et al. Malignant cells facilitate lung metastasis by bringing their own soil. Proc. Natl Acad. Sci. USA. 2010;107:21677–21682. doi: 10.1073/pnas.1016234107. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 27.Nakhasi HL, Grantham FH, Gullino PM. Expression of κ-casein in normal and neoplastic rat mammary gland is under the control of prolactin. J. Biol. Chem. 1984;259:14894–14898. [PubMed] [Google Scholar]
  • 28.Gullino PM. Tumor pathophysiology: the perfusion model. Antibiot. Chemother. 1980;28:35–42. doi: 10.1159/000386057. [DOI] [PubMed] [Google Scholar]
  • 29.Gullino PM, Grantham FH. Studies on the exchange of fluids between host and tumor. I. A method for growing ‘tissue-isolated’ tumors in laboratory animals. J. Natl. Cancer Inst. 1961;27:679–693. [PubMed] [Google Scholar]
  • 30.Kristensen CA, Roberge S, Jain RK. Effect of tumor necrosis factor alpha on vascular resistance, nitric oxide production, and glucose and oxygen consumption in perfused tissue-isolated human melanoma xenografts. Clin. Cancer Res. 1997;3:319–324. [PubMed] [Google Scholar]
  • 31.Bayless KJ, Kwak HI, Su SC. Investigating endothelial invasion and sprouting behavior in three-dimensional collagen matrices. Nat. Protoc. 2009;4:1888–1898. doi: 10.1038/nprot.2009.221. [DOI] [PubMed] [Google Scholar]
  • 32.Albini A, Benelli R. The chemoinvasion assay: a method to assess tumor and endothelial cell invasion and its modulation. Nat. Protoc. 2007;2:504–511. doi: 10.1038/nprot.2006.466. [DOI] [PubMed] [Google Scholar]
  • 33.Proia DA, Kuperwasser C. Reconstruction of human mammary tissue in a mouse model. Nat. Protoc. 2006;1:206–214. doi: 10.1038/nprot.2006.31. [DOI] [PubMed] [Google Scholar]
  • 34.Bockhorn M, Roberge S, Sousa C, Jain RK, Munn LL. Differential gene expression in metastasizing cells shed from kidney tumors. Cancer Res. 2004;64:2469–2473. doi: 10.1158/0008-5472.can-03-0256. [DOI] [PubMed] [Google Scholar]
  • 35.Swartz MA, et al. Cells shed from tumours show reduced clonogenicity, resistance to apoptosis, and in vivo tumorigenicity. Br. J. Cancer. 1999;81:756–759. doi: 10.1038/sj.bjc.6690760. [DOI] [PMC free article] [PubMed] [Google Scholar]

RESOURCES