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[Cancer Research 65, 8324-8330, September 15, 2005]
© 2005 American Association for Cancer Research

Experimental Therapeutics, Molecular Targets, and Chemical Biology

In vivo Assessment of RAS-Dependent Maintenance of Tumor Angiogenesis by Real-time Magnetic Resonance Imaging

Yi Tang1, Minjung Kim2, Daniel Carrasco2,5, Andrew L. Kung3, Lynda Chin2,4 and Ralph Weissleder1

1 Center for Molecular Imaging Research, Massachusetts General Hospital, Harvard Medical School, Charlestown, Massachusetts; Departments of 2 Medical Oncology and 3 Pediatric Oncology, Dana-Farber Cancer Institute and Children's Hospital; and Departments of 4 Dermatology and 5 Pathology, Brigham and Women's Hospital, Harvard Medical School, Boston, Massachusetts

Requests for reprints: Lynda Chin, Dana-Farber Cancer Institute, 44 Binney Street, M413, Boston, MA 02115. Phone: 617-632-6091; E-mail: lynda_chin{at}dfci.harvard.edu or Ralph Weissleder, Center for Molecular Imaging Research, Massachusetts General Hospital, Room 5406, Building 149, 13th Street, Charlestown, MA 02129. Phone: 617-726-8226; Fax: 617-726-5708; E-mail: weissleder{at}helix.mgh.harvard.edu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results and Discussions
 References
 
New blood vessel formation is a prominent feature of human cancers and tumor progression and is frequently accompanied by the acquisition of an angiogenic phenotype associated with a switch in the balance of proangiogenic and antiangiogenic molecules. This study was designed to investigate the role of activated H-RAS on the angiogenic phenotype of melanoma that arises in the inducible Tyr/Tet-RAS Ink4a/Arf–/– model using in vivo imaging with histopathologic correlation. We show that loss of RAS activity in fully established melanomas led to a reduction in tumor volume, which was preceded by impairment of vascular function as determined by in vivo magnetic resonance imaging. This correlated with activation of apoptosis in host-derived endothelial cells as well as in tumor cells. Thus, real-time in vivo imaging provided evidence that maintenance of tumor angiogenesis requires activated RAS in this model system, and that loss of vascular integrity upon inactivation of RAS is an active process rather than a consequence of loss of tumor cell viability.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results and Discussions
 References
 
A tumor represents the phenotypic end point of successive genetic lesions that alter the function and regulation of oncogenes and tumor suppressor genes. To achieve an immortal state, a normal cell must attain (a) unconstrained cell growth by disruption of the retinoblastoma pathway; (b) inactivation of a p53-dependent apoptotic response elicited by such inappropriate proliferation; and (c) maintenance of telomere function achieved most often by reactivation of telomerase. This trio of events constitutes the fundamental requirement that impart indefinite growth potential to cells in culture. However, the genetic roadmap beyond immortalization toward a fully transformed tumorigenic state is less well understood. In its fully transformed state, an immortal cell has acquired the ability to commandeer host functions to establish a permissive and supportive microenvironment in which a tumor can grow and survive (1). The resultant tumor is then maintained and sustained through complex and poorly understood processes and represents rational points of therapeutic intervention. Angiogenesis is generally considered as such a process that plays an essential role in delivering nutrients required for malignant tumor growth, invasion, and metastasis (1).

New blood vessel formation is a prominent feature of human cancers and tumor progression is frequently accompanied by the acquisition of an angiogenic phenotype, associated with a switch in the balance of proangiogenic and antiangiogenic molecules (2). Immunohistochemical quantification of intratumoral vessels stained for the vascular junction molecule CD31 (platelet/endothelial cell adhesion molecule 1) is commonly used as a marker of microvessel density. Similarly, factor VIII–related antigen has been used to detect tumor vessels, although this protein is only expressed by a fraction of immature, CD31-positive intratumoral blood vessels. Lectin Ulex europaeus agglutinin I has also been used for morphometric analysis of blood vessels (3). These methodologies are primarily applied ex vivo, typically requiring "hotspot" or representative field analysis, and are prone to missampling. Most importantly, these techniques do not permit the study of the role of angiogenesis in real time and serially in vivo.

A number of elegant intravital confocal or multiphoton approaches have recently been described to assess tumor neovasculature in intact animals (4, 5). Most of these approaches rely on transgenic models expressing targeted green fluorescent protein and/or the injection of a large molecular weight fluorescent marker, such as fluorescent dextrans, nanoparticles, or other preparations (4, 6). Whereas these methods have shed light onto microscopic detail of single vessels and angiogenic networks, they are generally not suitable for survey of an entire millimeter-sized tumor in three-dimensional fashion and longitudinally over time. We reasoned that analogous materials could be used for measuring tumor vascular parameters by high-resolution magnetic resonance imaging (MRI). One highly successful approach has been the use of long circulating (>10 hours) magnetic nanoparticles to probe microvascular changes in inflammation (7) and tumor environments (8). The nanoparticles contain a small, monocrystalline, magnetic iron oxide core, which exhibits strong magnetic behavior detectable by high-resolution MRI. The 3 nm core is surrounded by a dense, modified dextran coating that diminishes the immunogenicity of the assembled particles (20-30 nm) and substantially enhances their half-life in circulation. A further sophistication has been the generation of coatings modified with "tags," such as fluorochromes (magnetofluorescent nanoparticles) or radioisotopes, thereby permitting detection by additional imaging techniques (9). These vascular probes also have other attractions, such as their nontoxicity and promising performance in clinical trials. Indeed, MRI of related materials was recently applied with success to patients with prostate cancer, enabling visualization of tumor angiogenesis as well as small and otherwise undetectable lymph node metastases (10). Here, we apply magnetic nanoparticles to measure the vascular volume fraction in entire mouse tumors in real time (8).

Previously, we have generated and characterized an inducible RAS-driven melanoma model on the background of Ink4a/Arf deficiency (11). In this model, activated RAS expression is regulated by doxycycline in the media or in drinking water in vitro or in vivo, respectively (11). These mice develop spontaneous cutaneous melanomas in a strictly doxycycline-dependent manner. When RAS expression was down-regulated in Tyr/Tet-RAS Ink4a/Arf–/– mice bearing established melanomas, complete clinical regression ensures within 10 to 14 days. Thus, this model represents an ideal system in which to examine processes critical for tumor maintenance. For instance, we have observed that tumor regression upon loss of RAS activity is accompanied by activation of apoptosis affecting tumor and host-derived endothelial cells. This suggests an active role for RAS in sustaining tumor angiogenesis and interruption leads to vascular collapse during tumor regression. However, histopathologic studies were unable to ascertain the onset of endothelial cell apoptosis relative to overt tumor cell death. Therefore, the question remains whether endothelial cell death is merely a reactive process consequent to overall tumor cell death. To address this, we used real-time serial MRI to measure the kinetics of vascular impairment relative to tumor volumetrics measured by bioluminescence (cell activity) and metric methods (MRI three-dimensional volume, calipers).


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results and Discussions
 References
 
Melanoma cell line, transgenic mice, and severe combined immunodeficient mice explant. The inducible Tyr/Tet-RAS transgenic mice on a Ink4a/Arf-deficient background have been described (11, 12). Mice were fed with doxycycline in drinking water (2 mg/mL in sucrose water) to activate H-RasV12G expression in melanocytes. Tumors were allowed to develop over 6 to 10 weeks. Mice harboring single or multiple melanomas were imaged on day 0 and day 3 following doxycycline withdrawal and subsequent RAS down-regulation. Analyses were done on animals of FVB (N10) background.

R545 cells (11), melanoma cells derived from Tyr/Tet-RAS Ink4a/Arf–/– mouse, were implanted s.c. into both flanks of CB-17-scid (C.B-Igh-1b/IcrTac-Prkdcscid; Taconic, Germantown, NY) mice at 106 cells per site. Tumors were allowed to develop over 2 to 3 weeks under doxycycline (2 mg/mL in sucrose water) and were imaged serially at the indicated time points following down-regulation or reactivation of RAS expression. Specimens were collected following imaging for the histologic analyses.

For the retroviral transduction, retroviral vector (pbabe-puro-Luciferase) was transfected into 293T cells using the pCL-Eco helper plasmid. Retroviral supernatants isolated 36 and 60 hours after transfection were used to infect R545 cells. At 24 hours postinfection, the cells were selected for 2 days in growth medium containing 2 µg/mL puromycin. Cells were passaged no more than two passages before s.c. injection into severe combined immunodeficient (SCID) mice. R545-vascular endothelial growth factor (VEGF) cells were as described previously (11).

Histologic analysis and immunohistochemistry. Tissue samples were formalin fixed and paraffin embedded. Apoptosis was visualized by terminal deoxynucleotidyl transferase–mediated nick end labeling (TUNEL) assay (ApopTag kit; Chemicon, Temecula, CA), blood vessels were stained with anti-CD31 antibody (BD PharMingen, San Diego, CA), and proliferating cells were marked with Ki67 antibody (Novocastra Lab, Newcastle, United Kingdom). Immunodetection was done using a Vector Elite ABC kit and a Vector NovaRED kit for substrate detection. TUNEL-positive cells were counted from six random high power fields for each time points in a blind manner and normalized for the cell numbers per each field. Total vessel perimeter and proliferating cells were measured per high power field from five random fields in a blinded way. Differences between groups were analyzed by the unpaired t test with Welch's correction.

Magnetic resonance imaging. All MRI studies were carried out using a 1.5 T clinical MRI system (Signa; GE Medical Systems, Milwaukee, WI) and clinically available pulse sequences in anticipation that this technology would be directly applicable to a clinical setting. Tumor-bearing mice were anesthetized with ketamine (80 mg/kg i.p.) and xylazine (12 mg/kg i.p.). Custom-made 28-g catheters were inserted into a lateral tail vein and attached to a microheparin-saline flush unit. A 3-in. surface receiver coil was used for image acquisition. Following a fast spoiled gradient echo localizer sequence, multiple axial gradient-echo sequence were obtained (TR/TE 3,000/20, a 90-degree flip angle) using a 256 x 256 matrix and one excitation. The field of view was set at 10 x 4 cm, and the section thickness was 1.5 mm. All animals were imaged before and after i.v. injection of magnetic nanoparticles (5 mg of Fe per kilogram of MION-47; Center for Molecular Imaging Research, Massachusetts General Hospital, Boston, MA). Animals were kept warm by placing them on a heating pad during imaging. The entire imaging time was ~20 minutes for each mouse. At the end of serial MRI experiments of each group, the animals were sacrificed by means of ketamine and xylazine overdose. All animal studies were approved by the Institutional Animal Care Committee.

Steady-state tumoral blood volume maps were calculated from series of precontrast and postcontrast magnetic nanoparticle images, as described in detail elsewhere (8, 13). A fundamental observation is that the change in the transverse relaxation rate ({Delta}R2*) relative to the preinjection baseline relaxation rate is proportional to the perfused local blood volume per unit tumor volume (V) multiplied by a function (f) of the plasma concentration of the agent (P): {Delta}R2* = k x f(P) x V. This observation also forms the basis for neurofunctional MRI (14, 15). Following steady state of magnetic nanoparticle distribution (within minutes), the equation is revised to express a simple linear relationship between the change in the transverse relaxation rate and the perfused blood volume fraction: {Delta}R2*(t) = k x V(t) or V(t) = ({Delta}R2*(t)) / k, where {Delta}R2*(t) is the change in the transverse relaxation rate of the tumor, V(t) is the tumor volume, and the constant k includes the blood concentration and is, therefore, dose dependent. The enhancement of transverse relaxation can thus be expressed as follows: {Delta}R2* = ((1 / T2*post) – (1 / T2*pre)) {approx} (–1 / TE) (ln(Spost/Spre)), where S is the signal intensity; TE, the echo time; and T2*, the transverse relaxation time. On the basis of this formula, maps depicting the change in transverse relaxation rate were calculated from all the magnetic resonance images by using a homemade software (CMIR Image, Massachusetts General Hospital, Boston, MA). Absolute tumoral vascular volume fraction were obtained by scaling measurements to muscle with a known vascular volume fraction of 3% (16) and by having done previous calibration curves with a nuclear marker of plasma volume (8, 17). The major sources of error with MRI measurements occur by either animal movement during image acquisition (minimized by immobilization frame), temperature and anesthesia effects (minimized by use of fluothane and heating pads), and the method of gradient echo acquisition (pulse sequence) and subsequent T2* calculation (multiecho train measurements more accurate than two-echo measurements).

For each magnetic resonance slice and respective echo frames, regions of interest were drawn encompassing the entire tumor and separately adjacent muscle (20-100 pixels). Three-dimensional tumor volumes and vascular volume fraction (mean and maximum) were then calculated. To display the imaging results, the vascular volume fraction maps were superimposed onto anatomically coregistered magnetic resonance images. One-way ANOVA was used for statistical analysis.

Bioluminescence imaging. Bioluminescence imaging was done with a cryogenically cooled high-efficiency charge-coupled device (CCD) camera system (Xenogen IVIS 100). Mice were injected i.p. with D-luciferin (150 mg/g body weight; Biotium, Hayward, CA) and images were acquired 5 to 10 minutes after D-luciferin administration. Surface images of each animal were acquired under dim polychromatic illumination. Luciferase activity from the implanted R545 tumors was then measured by recording photon counts in the CCD with no illumination. Image postprocessing and visualization was done with a home-written program (CMIR Image, Massachusetts General Hospital) or the custom supplied software. Regions of interest were defined and recorded as mean, SD, and sum of the photon counts per unit time. For visualization purposes, the bioluminescence images were fused with the corresponding white light surface images as a pseudocolor overlay, permitting correlation of areas of bioluminescence activity with anatomy.


   Results and Discussions
Top
 Abstract
 Introduction
 Materials and Methods
 Results and Discussions
References
 
Vascular impairment precedes overall tumor reduction upon RAS inactivation. We had previously shown that melanoma cell lines (such as R545) derived from Tyr/Tet-RAS Ink4a/Arf–/– animals recapitulate the RAS-dependent tumor growth and regression phenotypes observed in de novo tumor-bearing transgenics (11). Thus, we first did a side-by-side time course comparison of in vivo MRI and standard histopathology in tumors derived from the congenic cell line R545. We injected the well-characterized R545 cells (11) into both flanks of 12 SCID mice, which were maintained on doxycycline drinking water until tumor growth reached ~0.5 cm in diameter. Upon doxycycline withdrawal, MRI with MION was done on three mice at each time point (on days 0, 1, 3, and 5 following RAS inactivation). After imaging, mice were sacrificed and tumors harvested for detailed histopathologic characterization.

As shown previously by caliper measurement, R545-derived SCID tumors did not exhibit apparent tumor shrinkage until day 5 (11), Ki67 immunostaining showed significant reduction in the proliferation index only on day 5 (day 0 versus day 5; P < 0.0001) although a moderate decrease was noted on day 3 (Fig. 1A and C). However, apoptosis involving both tumor cells and host-derived endothelial cells was activated on day 3, with TUNEL-positive cells per 1,000 nuclei increasing from 5.8 and 6.4 on day 0 and day 1, respectively, to 40.3 on day 3 and remaining high on day 5 (Fig. 1A and C). Consistent with apoptotic involvement of endothelial cells (Fig. 1C, inset), CD31 immunohistochemistry revealed significant reduction in vessel density as measured by vessel perimeters, which decreased from 86.5 ± 16.4 cm on day 0 to 54.7 ± 12.6 cm on day 5 (P = 0.0108, unpaired t test with Welch's correction; Fig. 1B). Although the difference in vessel perimeter counts was not statistically significant (day 0 versus day 3, P = 0.1318), vessel morphology had changed by day 3 and was characterized by fragmentation of CD31-positive "vessels," suggesting a collapse in tumor vasculature (Fig. 1C, compare Ci-Cl).



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  		Figure 1. Histologic analyses of R545 explant tumors during regression. Quantitation of apoptotic cells and proliferating cells (A), blood vessels (B), and representative images (C) are shown. TUNEL (Ce-Ch) shows activation of apoptosis from day (D) 3, and inset on (Cg) shows apoptosis of host-derived endothelial cell. CD31 (platelet/endothelial cell adhesion molecule; Ci-Cl) staining illustrates blood vessel collapse from day 3. Ki67 staining marks proliferating cells (Ca-Cd). Ca, Ce, Ci, day 0; Cb, Cf, Cj. day 1; Cc, Cg, Ck, day 3; and Cd, Ch, Cl, day 5 following RAS down-regulation.

 
In comparison to the ex vivo histopathologic analysis, in vivo MRI detected significant vascular impairment by day 3. Both the mean and maximum vascular volume fractions decreased dramatically from day 0 to day 3 (P = 0.001; Fig. 2). Moreover, a decrease in vascular volume fraction was observed within 24 hours in some animals (data not shown), suggesting that MRI with MION was more sensitive than CD31 immunohistochemistry in detecting impairment in vascular function. Consistent with caliper measurements, the mean tumor volumes steadily decreased over time (MRI measurements: 521 ± 91 mm3 on day 1, 483 ± 216 mm3 on day 3, and 110 ± 22 mm3 on day 5). This decrease was only statistically significant on day 5 (P < 0.05 compared with day 0), lagging behind the reduction in vascular volume fraction. Comparison of these two parameters in matched pairs through regression analysis showed that no significant correlation existed between tumor size and vascular volume fraction (r2 = 0.04, P = 0.5; data not shown). Furthermore, this lag in tumor volume reduction was not an artifact reflecting delay in elimination of dead tumor cells by host macrophages, because the tumors on day 3 were viable as shown by Ki67 staining (Fig. 1A and C) and by bioluminescence imaging of cell activity (Fig. 2D).



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  		Figure 2. Temporal changes of tumor vascular volume fraction and tumor volume following down-regulation of RAS. A and B, both mean and maximum vascular volume fraction (VVF) decrease significantly after 3 days of RAS inactivation, whereas the tumor volume decreases later (day 5), suggesting that vascular impairment is the preceding event. C, color-coded vascular volume fraction tumoral vascularity maps derived from precontrast and postcontrast T2*-weighted MRI are superimposed onto the tumors. Note the heterogeneity of vascular volume fraction among the tumors. D, tumor mass measured by bioluminescence (day 0 and day 3 only) and calipers upon doxycycline withdrawal. Histogram summarizes the time course of four mice. Mean photon flux on day 0 was 2.5 ± 2.3 (photons/s) and 4.3 ± 3.4 on day 3. Note robust cell activity by bioluminescence on day 3, ruling out possibility of dead tumor cells despite lack of tumor volume decrease until day 5 (by caliper measurement).

 
Finally, we asked whether enforced expression of VEGF could reverse the loss of vascular volume fraction after RAS inactivation by utilizing R545 cells engineered to express high levels of VEGF by retroviral infection (R545-VEGF; ref. 11). Here, SCID mice bearing R545-VEGF–derived explant tumors on doxycycline were imaged serially by MRI at baseline (day 0) and 5 days after doxycycline withdrawal. As shown in Fig. 3, enforced VEGF expression was not able to reverse the collapse in vascular function after RAS inactivation, as significant decreases for mean and maximum vascular volume fraction on day 5 were observed (P = 0.0153 and P = 0.0075, respectively). This is consistent with our previous finding that VEGF expression was not able to rescue endothelial cell viability after RAS inactivation in vivo (11), suggesting that other mediators, in addition to VEGF, are important in mediating RAS-dependent maintenance of tumor vascularity.



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  		Figure 3. Temporal changes of tumor volume and vascular volume fraction in R545-VEGF model. Note decrease in tumor volume, as well as vascular volume fraction, by day 5 after doxycycline withdrawal despite enforced VEGF expression.

 
In summary, MRI of explant tumors derived from the R545 congenic cells revealed that vascular function impairment was an early consequence of RAS inactivation, preceding gross loss in tumor volume. This observation suggests that maintenance of endothelial cell viability and tumor vasculature is an active process dependent on RAS activities that extend beyond VEGF regulation.

RAS plays a role in maintenance of tumor vascular function. To more directly address whether RAS activity maintains vascular integrity in an established tumor, we did serial imaging of the same tumor-bearing mice during RAS inactivation and reactivation (see Fig. 4A for experimental design). Following baseline MRI on day 0, tumor-bearing mice were randomly assigned to either a control cohort (maintained on doxycycline with continued RAS activity, n = 4) or an experimental cohort (subjected to 3 days of RAS inactivation followed by 5 days of RAS reactivation, n = 4). Tumor volumes between the two groups were similar on day 0 (598 ± 267 versus 557 ± 315 mm3; Fig. 4D). Serial imaging was done on day 3 and day 8 for all animals in both cohorts.



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  		Figure 4. Serial analyses of tumor size and vascular volume fraction after RAS inactivation and reactivation. A, summary of the experimental design. A set of representative images is shown in (B-C) with color-coded tumoral vascularity. Note the decrease in vascular volume fraction on day 3 in the experimental group with partial recovery by day 8. D, summary of the tumoral volume changes following activation and inactivation of RAS. Significant difference between the experimental and control group was noted on day 8. Although RAS was reactivated for 5 days in the experimental group, the shrinkage effect caused by the preceding RAS inactivation contributes to the significant differences on day 8. E, histopathology of explant tumors on day 8 following final MRI. TUNEL and CD31 staining shows suppression of cell death and restoration of vessels in experimental group, which makes them comparable with the control group on day 8.

 
Three days after RAS inactivation, there was a significant loss in vascular volume fraction in the experimental animals compared with the control (43.8 ± 12.4 versus 99.6 ± 17.9, P < 0.001; Fig. 4B-C), whereas tumor volumes remained largely unchanged (P > 0.05; Fig. 4D). Interestingly, after 5 days of RAS reactivation (day 8 imaging), the vascular volume fraction improved, but did not recover completely when compared with the control (77.2 ± 13.4 versus 99.8 ± 18.8, P < 0.05; Fig. 4C). Concomitant with this improvement in vascular function, CD31 immunohistochemistry revealed restoration of intact vascular lumens in day 8 specimens (total vessel perimeters: 125.9 ± 28.0 cm in the control group versus 123.3 ± 11.6 cm in the experimental group, P = 0.8452; Fig. 4E). Similarly, apoptosis measurements revealed a dramatic suppression of cell death upon reactivation of RAS, with 5.0 TUNEL-positive nuclei per 1,000 on day 8 versus 4.8 on day 0 (P = 0.8985; Fig. 4E). Thus, this study shows that the tumor vascular function requires RAS activities and that continuous RAS activity in tumor cells plays a role in maintaining intact functional tumor vasculature.

To further show that the in vivo vascular kinetics observed by serial MRI was not unique to the specific congenic cell line used in the explant studies, we did a similar serial imaging study in transgenic Tyr/Tet-RAS Ink4a/Arf–/– animals with established de novo tumors (see Fig. 5A for experimental design). Six cutaneous melanomas from two different transgenic mice (three tumors per mouse) were imaged serially on day 0 and day 3 following RAS down-regulation for tumor volume and vascular volume fraction by MRI. As before, whereas alteration in tumor volume was not statistically significant (paired t test, P = 0.3462), vascular volume fraction was significantly reduced (paired t test, P = 0.0132 for mean vascular volume fraction and P = 0.0058 for maximum vascular volume fraction; Fig. 5B-C). Importantly, all six tumors showed reduced mean and maximum vascular volume fraction on day 3 compared on day 0, ranging from 11.7% to 66.5% for mean vascular volume fraction and 17.2% to 39.4% for maximum vascular volume fraction (Fig. 5C; Table 1). Interestingly, although one tumor showed continued volume increase on day 3 (Table 1, mouse 2, tumor 6), vascular volume fraction reduction was evident. Finally, it is worth noting that, in contrast to the explant studies with the congenic cell line, this de novo tumor study revealed a much wider variation in vascular volume fraction kinetics. This is not unexpected because complex tumor phenomenon, such as angiogenesis, are regulated on multiple levels by multiple genetic lesions, thus dependence of RAS activity will vary in the context of other mutations. Such genetic heterogeneity of independent de novo tumors arising in engineered models is a more accurate reflection of the human condition.



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  		Figure 5. Temporal changes of vascular volume fraction and tumor volume of de novo tumors after down-regulation of RAS expression. Serial imaging of two Tyr-rtTA/Tet-RAS Ink4a/Arf–/– transgenic mice bearing six independent cutaneous melanomas on day 0 and day 3 after doxycycline withdrawal. Experimental design is outlined in (A). All six de novo tumors showed significant decreases in both mean and maximum vascular volume fraction after 3 days of Ras inactivation, and five of the tumors decreased in volume (B; see also Table 1). Color-coded vascular volume fraction tumoral vascularity maps derived from precontrast and postcontrast T2*-weighted MRI are superimposed onto anatomic images. Note the shrinkage of tumor volume and decrease of vascular volume fraction of the de novo tumor (C).

 

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  		Table 1. Percentage decrease in tumor volumes and vascular volume fractions on day 3 compared with day 0 after doxycycline withdrawal

 
In conclusion, our studies presented here show that real-time MRI measurements of vascular volume fraction are a reliable biomarker of tumor vascular function compared with traditional ex vivo immunohistopathologic characterization. It offers enhanced sensitivity in capturing impairment in vascular function before apparent morphologic change in tumor vasculature. Moreover, real-time measurement of vascular function provides strong evidence that the maintenance of tumor angiogenesis requires activated RAS in this model. Upon inactivation of RAS, loss of vascular integrity is an active process rather than a consequence of loss of tumor cell viability.


   Acknowledgments
 
Grant support: Grants R24 CA92782 (Small Animal Imaging Resource Program) and P50 CA86355 (In vivo Cellular and Molecular Imaging Center; R. Weissleder); R01 CA93947 and U01 CA84313 (L. Chin); and 1K 08AG01031-02 (D. Carrasco).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Renee D. Wright for technical assistance with the bioluminescence studies and Dasha Chestukhin for assistance with data quantitation.


   Footnotes
 
Note: Y. Tang and M. Kim contributed equally to this work. L. Chin is a Charles E. Culpepper Scholar.

Received 1/ 5/05. Revised 7/ 7/05. Accepted 7/14/05.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results and Discussions
 References
 

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