786-O cells (VHL deficient human renal cell carcinoma) were obtained from the American Type Culture Collection (ATCC, Manassas, VA) and were grown in RPMI 1640 medium from Cellgro. All media were supplemented with 2 mM L-glutamine, 10% fetal calf serum and 1% streptomycin (50 μg/ml) and cells were cultured at 37°C with 5% CO2.
Tumor xenograft induction
For the subcutaneous xenograft tumor model female nude beige mice (Charles River Laboratories, MA), 6-8 weeks of age and 20 g average body weight, were used as per . The mice were housed and maintained in laminar flow cabinets under specific pathogen-free conditions. All experiments were approved by the Institutional Animal Care and Use Committee at Beth Israel Deaconess Medical Center. To produce tumors, renal cancer cells were harvested from subconfluent cultures by a brief exposure to 0.25% trypsin and 0.02% EDTA. Trypsinization was stopped with medium containing 10% FBS, and the cells were washed once in serum-free medium and resuspended in PBS. Only suspensions consisting of single cells with greater than 90% viability were used for the injections.
To establish RCC tumor xenograft, an established human VHL deficient RCC cell line (786-O) was injected subcutaneously (1 × 107 cells) into the flanks of 6-8 week old nude/beige mice. Tumors developed in > 80 percent of the mice and were usually visible within 2 weeks of implantation and once they reached a diameter of 3-5 mm, were measured daily to ensure a consistent size at the outset of treatment. Tumor long and short axes were measured using calipers daily. Tumor volumes were calculated with the formula volume = length × width2/2 and followed to determine growth curves. Animals were euthanized according to IACUC guidelines and treatment was terminated as experimentally designed and described below. Prior to dissecting the tumor, the midpoint of the cranial caudal axis of the tumor was marked. This marked line matched the ASL imaging slice. The tumor was cut into three equal segments parallel to the marked line. The mid segment of the tumor was fixed in 10% formalin at room temperature for 24 hours prior to embedding in paraffin. Tumors were sectioned and stained with H&E, and immunohistochemical analysis.
Sorafenib tosylate (80 mg/kg daily 6 of 7 days per week by gavage) was begun when the tumors had grown to a diameter of 12 mm [14, 15]. This 80 mg/kg dose was based on a study by Chang et al.  in which four doses of sorafenib (15, 30, 60 and 90 mg/kg; free base equivalent) were compared in 786-O and RENCA xenograft mice. They demonstrated similarity in the 60 and 90 mg/kg groups in terms of growth delay. The 60 mg/kg free base dose, would convert to 82 mg/kg of the tosylate form that was used in this study. This dose (rounded to 80 mg/kg) was considered the maximally effective dose. It was used in our prior studies with this murine model  and was defined as the "conventional dose" for current study. The "high dose intermittent" regimen was 160 mg/kg administered 3 days on and 4 days off, "low dose intermittent" was 80 mg/kg administered 3 days on and 4 days off, and "high dose continuous" therapy was 160 mg/kg administered continuously. The continuous conventional dose and the high dose intermittent regimens delivered the same total dose of sorafenib over 7 days (80 mg/kg/day given 6 out of 7 days per week and 160 mg/kg/day 3 days per week).
Mice were grouped randomly into treatment with vehicle (n = 9), conventional dose continuous (n = 11), high dose intermittent (n = 11), conventional dose intermittent (n = 8), and high dose continuous (n = 4) by gavage when the tumors reached 12 mm in diameter. All animals were sacrificed and tumors were dissected ~36 days post therapy with the exception of vehicle treated mice which were sacrificed when tumors reached the mandated 20 mm sacrifice size (~22 days post treatment). In addition, we sacrificed 6 mice on day 3 after treatment with high dose (n = 3) and conventional dose (n = 3) for both the CD34 and CD31 analyses.
ASL MR imaging was performed on 6 mice prior at baseline (day 0), day 3, day 7, and day 10 post treatment with conventional dose continuous (n = 3) or high dose intermittent (n = 3).
For CD34 analysis, 4 um thick sections were prepared from formalin-fixed, paraffin-embedded tumor specimens. Sections were deparaffinized, rehydrated and heated with a pressure cooker to 125°C for 30 seconds in citrate buffer for antigen retrieval. After cooling to room temperature, sections were incubated in 3% hydrogen peroxide for 5 minutes to quench endogenous peroxidase, (Dako, Carpinteria, CA). The anti-CD34 antibody (Abcam, Cambridge, MA, Cat # AB-8158) was applied at a 1:50 dilution, diluted with DaVinvi Green diluent (BioCare Inc, Cat# PD900L), to sections for 1 hour, followed by rabbit anti-rat secondary antibody for 30 minutes. Detection was performed by incubating with Dako EnVision+ System HRP labeled polymer anti-rabbit for 30 minutes, followed by DAB chromogen. Slides were scanned using the Scanscope XT (Aperio Technologies Inc., Visa, CA) and analyzed using a modified Microvessel analysis algorithm (Aperio Technologies Inc).
For CD31 analysis, 4 um thick sections were prepared from formalin-fixed, paraffin-embedded tumor specimens. Sections were deparaffinized, rehydrated and heated with a pressure cooker to 125°C for 30 seconds in citrate buffer for antigen retrieval. After cooling to room temperature, sections were incubated in 3% hydrogen peroxide for 5 minutes to quench endogenous peroxidase, (Dako, Carpinteria, CA). The anti-CD31 antibody (Abcam, Cambridge, MA, Cat #AB28364) was applied at a 1:50 dilution, diluted with Dako antibody diluent (Dako, Cat #S0809), to sections for 1 hour. Detection was performed by incubating with Dako EnVision+ System HRP labeled polymer anti-rabbit for 30 minutes, followed by DAB chromogen. Slides were scanned using the Scanscope XT (Aperio Technologies Inc., Visa, CA) and analyzed using a modified Microvessel analysis algorithm (Aperio Technologies Inc).
Tumor perfusion imaging
Tumor perfusion imaging (ASL MRI) was performed as previously described . Briefly mice were anaesthetized, and placed in the supine position on a 3 cm in diameter custom-built surface coil. Adhesive tape was used to limit movement. Images were acquired using a 3.0 T whole-body clinical MRI scanner (3T HD; GE Healthcare Technologies, Waukesha, WI). A single slice ASL image was obtained with a single-short fast spin echo sequence (SSFSE) using a background-suppressed, flow-sensitive alternating inversion-recovery strategy. Twenty-four label and control pair images were acquired and averaged for the ASL acquisition. A reference proton density image was acquired by turning off all background suppression and labelling pulses in the ASL preparation. T1 measurement was performed after ASL imaging by using the same imaging sequence at same slice location but with inversion recovery at different inversion times. The single transverse slice of ASL was carefully positioned at the center of the tumor, which was marked on the skin with a permanent marker pen for follow-up MRI studies. ASL sequence raw data were saved and transferred to the analysis workstation for image reconstruction by using custom software written within the Interactive Data Language (IDL; ITT visual Information Solutions, Boulder, CO). The ASL difference image, between average label and control images, was then converted to quantitative tumor perfusion as previously described .
Perfusion was calculated on a pixel-by-pixel basis, and quantitative maps were produced. The quantitative maps and the corresponding proton density reference images were then analyzed by using Image J software (Image Processing and Analysis in Java; National Institutes of Health, Bethesda, MD). To determine tumor perfusion, a region of interest was drawn freehand around the peripheral margin of the tumor by using an electronic cursor on the reference image that was then copied to the perfusion image. The mean blood flow for the tumor tissue within the region of interest was derived, and image window and level were fixed. A 16-color table was applied in 10 mL/100 g/min increments ranging from 0 to 160 mL/100 g/min, with flow values represented as varying shades of black, blue, green, yellow, red, and purple, in order of increasing perfusion.
All data were expressed as mean ± SEM. Statistical significance of differences among groups of sorafenib dosing response was calculated using one-way or two-way analysis of variance (ANOVA). Tukey's significant difference post hoc test was used for pairwise comparisons after ANOVA to correct for multiple testing. The groups among which P < 0.05 were considered significantly different.