SEARCH

SEARCH BY CITATION

Keywords:

  • photodynamic therapy (PDT);
  • verteporfin;
  • vascular targeting;
  • fluorescence imaging;
  • vascular permeability;
  • tumor perfusion;
  • enhanced green fluorescence protein (EGFP);
  • prostate tumor model

Abstract

  1. Top of page
  2. Abstract
  3. Material and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Photodynamic therapy (PDT) is a light-based cancer treatment modality. Here we employed both in vivo and ex vivo fluorescence imaging to visualize vascular response and tumor cell survival after verteporfin-mediated PDT designed to target tumor vasculature. EGFP-MatLyLu prostate tumor cells, transduced with EGFP using lentivirus vectors, were implanted in athymic nude mice. Immediately after PDT with different doses of verteporfin, tumor-bearing animals were injected with a fluorochrome-labeled albumin. The extravasation of fluorescent albumin along with tumor EGFP fluorescence was monitored noninvasively with a whole-body fluorescence imaging system. Ex vivo fluorescence microscopy was performed on frozen sections of tumor tissues taken at different times after treatment. Both in vivo and ex vivo imaging demonstrated that vascular-targeting PDT with verteporfin significantly increased the extravasation of fluorochrome-labeled albumin in the tumor tissue, especially in the tumor periphery. Although PDT induced substantial vascular shutdown in interior blood vessels, some peripheral tumor vessels were able to maintain perfusion function up to 24 hr after treatment. As a result, viable tumor cells were typically detected in the tumor periphery in spite of extensive tumor cell death. Our results demonstrate that vascular-targeting PDT with verteporfin causes a dose- and time-dependent increase in vascular permeability and decrease in blood perfusion. However, compared to the interior blood vessels, peripheral tumor blood vessels were found less sensitive to PDT-induced vascular shutdown, which was associated with subsequent tumor recurrence in the tumor periphery. © 2008 Wiley-Liss, Inc.

Photodynamic therapy (PDT) induces tumor destruction through a photochemical reaction involving a photosensitizer, light of a specific wavelength matching the absorption of the photosensitizer and molecular oxygen.1 Singlet oxygen, a product of this photochemical reaction, causes oxidative damage to target cells and tissues and is the primary reactive oxygen species responsible for the biological effects of PDT.2 Although direct tumor cyotoxicity and immune responses are involved as well, damage to the tumor vasculature has been shown to contribute significantly to the overall PDT effect of most photosensitizers.3

Verteporfin (the lipid-formulation of benzoporphyrin derivative monoacid ring A) is a photosensitizer that is currently approved for the treatment of age-related macular degeneration (AMD).4 We have shown previously that the dynamic distribution of verteporfin is predominantly intravascular at 15 min after intravenous injection and becomes mainly extravascular at 3 hr after injection.5 Based on this pharmacokinetic property, preferential tumor vascular targeting can be achieved by illumination at 15 min after verteporfin administration. We have been exploring this passive vascular targeting principle for the treatment of prostate tumors. Intravital fluorescence microscopy studies in the MatLyLu rat prostate tumor model has demonstrated that vascular-targeting PDT with verteporfin induces vascular permeability increase and thrombus formation, which ends in vascular shutdown and tumor necrosis.6 These results indicate that vascular-targeting PDT using verteporfin can be used for the management of localized prostate cancer.

Because vascular damage is the dominant effect of PDT, especially in the case of vascular-targeting PDT, it is important to study in detail how photosensitization modifies vascular functions. Most studies on PDT-induced tumor vascular changes have been done on excised tumor specimens after sacrificing the animals. Although they have been valuable in revealing microscopic details, such studies are only able to provide snap-shot information on each individual animal. To obtain longitudinal information in a single animal, noninvasive imaging techniques are necessary to examine vessel functional changes after PDT. Imaging modalities such as laser Doppler perfusion imaging,7, 8 diffuse correlation spectroscopy,9 laser speckle imaging,10, 11 optical coherence tomography12 and ultrasonography13 have all been shown to be useful techniques for monitoring tumor blood flow dynamics noninvasively after PDT. Moreover, noninvasive imaging using contrast agents allows one to follow perfusion changes and also provides real-time information regarding vascular permeability. For instance, angiography with fluorescent dyes such as fluorescein or indocyanine green is routinely used to examine vessel leakage and occlusion in AMD patients treated with PDT.14 Changes in tumor perfusion and vascular permeability after PDT have also been studied with contrast-enhanced MRI.15, 16

Because of its high sensitivity and versatility, in vivo fluorescence imaging is able to provide both macroscopic and microscopic longitudinal data in individual animals, which cannot be obtained in other ways.17–19 In this study, we used an in vivo whole-body fluorescence imaging system to monitor vascular response and tumor cell survival in an EGFP-expressing prostate tumor model following treatment with verteporfin-PDT. Moreover, we compared the in vivo tumor imaging results with the ex vivo fluorescence microscopy of frozen tumor sections. Our results indicate that the vascular response to vascular-targeting PDT is clearly different between tumor interior vessels and peripheral blood vessels.

Material and methods

  1. Top of page
  2. Abstract
  3. Material and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

Production and titer of lentivirus

Lentiviral production was performed as previously described.20 Briefly, we cotransfected pWPT-EGFP and third-generation packaging vectors into 293FT cells (Invitrogen Life Technologies) and collected culture supernatants after 48 and 72 hr of incubation in a 37°C and 5% CO2 incubator. We recovered virus by ultracentrifugation (1.5 hr at 25,000 rpm) in a Beckman SW28 rotor and resuspended the virus pellet in 25 μl of Opt-MEM media (Invitrogen Life Technologies). Viral titers were determined by infecting 293FT cells with serial dilutions of concentrated lentivirus followed by flow cytometry analysis 48 hr later. Typical viral preparations yielded 5 × 108 transducing units/ml.

Tumor cells and lentiviral transduction

R3327-MatLyLu rat prostate cancer cells were maintained in the RPMI-1640 medium with glutamine (Mediatech, Herndon, VA) supplemented with 10% fetal bovine serum (HyClone, Logan, UT) and 100 units/ml penicillin–streptomycin (Mediatech) at 37°C in a 5% CO2 incubator. For lentiviral transduction, the MatLyLu cells were infected with a multiplicity of infection of 50 and allowed to incubate overnight. Polybrene (8 μg/ml, Sigma) was used to facilitate lentiviral transduction. Supernatant was then removed after infection and replaced with complete RPMI-1640 growth medium. EGFP-transduced MatLyLu cells were examined with a fluorescence microscope at 48 hr after transduction. EGFP-MatLyLu cells were harvested, serial diluted and seeded in a 96-well plate with cell density of 1 cell per well. After incubation for 7 days at 37°C and 5% CO2 atmosphere, the clone exhibiting the highest EGFP fluorescence intensity was selected and expanded for subsequent experiments.

Animals and tumor models

Male NCr athymic nude mice (4–5 weeks old, National Cancer Institute, Frederick, MD) were used throughout the study. Tumors were induced by subcutaneous injection of about 1 × 105 EGFP-MatLyLu tumor cells in the thigh region of mice. Tumors were used for experiments when they reached a size of 5–7 mm in diameter. All animal procedures were carried out according to a protocol approved by the Institutional Animal Care and Use Committee (IACUC).

Photosensitizer

Verteporfin (benzoporphyrin derivative (BPD) in a lipid-formulation) was obtained from QLT (Vancouver, Canada) as a gift. A stock saline solution of verteporfin was reconstituted according to the manufacturer's instructions and stored at 4°C in the dark.

PDT treatments

A diode laser system (High Power Devices, North Brunswick, NJ) at 690-nm wavelength was used for the irradiation of EGFP-MatLyLu tumors. The laser was coupled to an optical fiber with 600 μm core diameter and expanded to generate an 11-mm diameter illumination spot through a collimator. Animals were anesthetized with injection (i.p.) of a mixture of ketamine (90 mg/kg) and xylazine (9 mg/kg) and tumors were exposed to light with an irradiance of 50 mW/cm2. Light intensity was measured with an optical power meter (Thorlabs, North Newton, NJ). Verteporfin was injected (i.v.) 15 min prior to light irradiation at a dose of 0.25 mg/kg.

Noninvasive tumor fluorescence imaging and image analysis

Tumor-bearing animals were i.v. injected with 20 mg/kg albumin labeled with tetramethylrhodamine isothiocyanate (TRITC-albumin, Sigma) immediately after PDT. EGFP-MatLyLu tumors were imaged with a noninvasive whole body fluorescence imaging system for the EGFP and TRITC signal before and at various times after treatment. The setup of this home-built broad beam imaging system has been described in detail in our previous paper.21 Briefly, the system includes a filtered white light source for excitation and a SensiCamQE high performance digital CCD camera (The Cook Corp, Auburn Hills, MI) to capture fluorescence emission passing through an emission filter. We used a 470/20 nm excitation filter and a 520/20 nm emission filter for imaging tumor EGFP fluorescence and a 535/20 nm excitation filter and 590-nm long-pass emission filter for imaging the TRITC fluorescence. Camera settings were kept constant for the control and PDT-treated animals throughout the imaging process. Animals were anesthetized by inhalation of 1.5% isofluorane and imaged first for EGFP and then TRITC fluorescence without moving the animals. The EGFP and TRITC images were pseudocolored and superimposed to generate composite images.

A 2.5-mm diameter region of interest (ROI) was centered over tumor or tumor-adjacent normal tissue areas, and the average EGFP and TRITC fluorescence intensities in the ROI were quantified with NIH ImageJ software. The fluorescence intensity in tumor or tumor-adjacent tissues after PDT was normalized to its own pretreatment value in each animal, and the data from different animals in each group were pooled to generate response curves. To determine the TRITC-albumin distribution in relation to tumor EGFP fluorescence, a straight line was drawn through the tumor tissue on composite images and the corresponding green (EGFP) and red (TRITC) intensities were measured along the line.

Tumor tissue fluorescence microscopy

Tumor-bearing animals were i.v. injected with 20 mg/kg Hoechst (Sigma) as a vascular perfusion marker at different time points after treatment. Animals were euthanized within 1 min after injection and tumor tissues were excised and snap-frozen in isopentane precooled with liquid nitrogen. Frozen tumor sections with thickness of 10 μm were cut and examined under a Leica DMI6000B fluorescence microscope with appropriate filter sets for Hoechst (excitation: 360/40 nm; emission: 470/40 nm) and TRITC (excitation: 546/12 nm; emission: 600/40 nm).

Tumor volume measurement and tumor histology

Three-dimensional tumor sizes were measured regularly after treatment by caliper, and the tumor volume was calculated using the formula π/6 × tumor length × tumor width × tumor height. Animals were euthanized at various time points after treatment. Tumor tissues were excised and fixed in 4% formalin solution. Fixed tumor tissues were dehydrated and then embedded in paraffin. Tissue sections with thickness of 5 μm were cut and stained with H&E.

Statistical analysis

Students' 2-tailed t-test was used to calculate statistical differences between 2 groups and the significance was accepted at p < 0.05. Statistical analysis was carried out using GraphPad software (GraphPad, San Diego, CA).

Results

  1. Top of page
  2. Abstract
  3. Material and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

The extravasation of TRITC-albumin, as indicated by the increase in TRITC fluorescence, was imaged noninvasively with a whole-body fluorescence imaging system. Figure 1 shows the TRITC fluorescence images (red) merged with tumor EGFP fluorescence images (green) at different time points after vascular-targeting PDT with verteporfin. PDT caused an overall increase in the TRITC fluorescence and this was more pronounced in the peritumor area. PDT-induced TRITC-albumin extravasation appeared to be dose dependent because the 50 J/cm2 light dose PDT caused a greater increase in the TRITC fluorescence compared to the 25 J/cm2 light dose treatment.

thumbnail image

Figure 1. In vivo fluorescence images of the TRITC-albumin extravasation and tumor EGFP fluorescence. The EGFP-MatLyLu tumors were illuminated with 25 or 50 J/cm2 light at 15 min after i.v. injection of 0.25 mg/kg dose of verteporfin. Immediately after treatment, tumor-bearing animals were i.v. injected with 20 mg/kg TRITC-albumin and imaged at different times after injection with a whole-body fluorescence imaging system as described in the materials and methods. Control tumors received no treatment. The images shown are the merged image of TRITC (red) and EGFP (green) fluorescence images. [Color figure can be viewed in the online issue, which is available at www.interscience.wiley.com.]

Download figure to PowerPoint

The average TRITC fluorescence intensity in tumor and tumor-adjacent normal tissue ROIs was quantified with NIH ImageJ software. It was observed that the average TRITC fluorescence in tumor areas was about 20% lower than tumor-adjacent normal tissue areas presumably because higher blood volume in tumor tissues causes more TRITC fluorescence quenching than in normal tissues.19 Both 25 and 50 J/cm2 PDT treatments significantly increased the TRITC fluorescence intensity in tumor (Fig. 2a, p < 0.05) and tumor-adjacent tissues (Fig. 2b, p < 0.05). Fluorescence intensity increase started from 1-hr post-PDT treatments and reached a plateau at about 4 hr thereafter while untreated control tumors exhibited little change in fluorescence intensity over the same period of time. In both tumor and tumor-adjacent tissues, PDT with 50 J/cm2 light dose induced a greater increase in the TRITC fluorescence intensity than the 25 J/cm2 light dose (p < 0.01). The 25 J/cm2 light dose PDT caused a similar increase (maximally about 1.5-fold increase) in the TRITC fluorescence intensity in both tumor and tumor-adjacent tissues (p > 0.05). The 50 J/cm2 PDT caused significantly higher TRITC fluorescence increase in tumor-adjacent tissues (about 3-fold increase at peak) compared to tumor tissues (about 2-fold increase at peak, p < 0.05).

thumbnail image

Figure 2. In vivo fluorescence image analysis showing (a) changes of the TRITC-albumin fluorescence intensity in tumor tissues, (b) changes of the TRITC-albumin fluorescence intensity in tumor-adjacent tissues, and (c) changes of tumor EGFP fluorescence intensity after treatment. The EGFP-MatLyLu tumors were treated with vascular-targeting PDT and imaged with a whole-body fluorescence imaging system. The TRITC and EGFP fluorescence intensities were measured in a circular 2.5 mm diameter ROI placed over the tumor or tumor-adjacent area on the fluorescence images. The fluorescence intensity values after treatment in each animal were normalized to their own pretreatment values, which are displayed as 100% at 0 time point. Each group included 3 or 4 animals. Error bars represent the standard deviation.

Download figure to PowerPoint

Changes in the average EGFP fluorescence intensity in tumor tissues were also quantified and are shown in Figure 2c. Both 25 and 50 J/cm2 PDT treatments caused a significant decrease in tumor EGFP fluorescence at 1 hr after treatment (p < 0.05). After the initial decrease, there was no further decrease in tumor EGFP fluorescence intensity. Control tumors showed little change in the EGFP fluorescence during this 5-hr period.

Analysis of TRITC and corresponding EGFP intensity profiles indicated that the TRITC fluorescence intensity in tumor peripheral area was higher than in tumor interior area at 4 hr after injection of TRITC-albumin (Fig. 3). However, an opposite pattern was found in tumor EGFP intensity profiles with the higher intensity values detected in the tumor center. Both 25 and 50 J/cm2 PDT treatments caused an overall increase in the TRITC intensity and decrease in tumor EGFP intensity. The increase in the TRITC intensity was found to be higher in the tumor periphery than in the tumor center.

thumbnail image

Figure 3. In vivo fluorescence image analysis showing the TRITC-albumin accumulation in relation to tumor EGFP fluorescence intensity. The 4-hr-time-point images from Figure 1 were analyzed and shown here. A 17-mm line was drawn through the tumor tissue on each fluorescence image. Both TRITC-albumin and tumor EGFP fluorescence intensities were measured along the line and are shown in the figure. Dashed lines indicate the boundary of the tumor tissue. Note the opposite pattern between tumor TRITC-albumin accumulation and tumor EGFP fluorescence intensity profiles. [Color figure can be viewed in the online issue, which is available at www.interscience.wiley.com.]

Download figure to PowerPoint

To verify the whole-body fluorescence imaging results, we euthanized animals at 1, 4 and 24 hr after 50 J/cm2 PDT treatment and excised tumor tissues for fluorescence microscopy. Hoechst dye was i.v. injected shortly before euthanizing animals to highlight functional blood vessels. As shown in Figure 4, tumor staining of Hoechst dye decreased significantly after vascular-targeting PDT with 50 J/cm2 light dose compared to the control tumor, indicating a decrease in functional blood vessels. Moreover, functional blood vessels were mainly detected at the tumor periphery after PDT. In agreement with the macroscopic in vivo tumor imaging results, fluorescence microscopy also demonstrated a significant increase in the TRITC fluorescence intensity after PDT, especially in the tumor periphery.

thumbnail image

Figure 4. Ex vivo fluorescence microscopy images showing the distribution of TRITC-albumin in relation to the functional blood vessels highlighted by Hoechst dye staining. The EGFP-MatLyLu tumors were treated with 50 J/cm2 light at 15 min after i.v. injection of 0.25 mg/kg dose of verteporfin. Control tumors received no treatment. Immediately after treatment, tumor-bearing animals were i.v. injected with 20 mg/kg TRITC-albumin. Animals were euthanized at 1, 4 or 24 hr after injection of the TRITC-albumin. Hoechst dye (20 mg/kg) was i.v. injected at 1 min before euthanizing the animal. Frozen tumor sections from tissue samples were first imaged for Hoechst fluorescence and the same fields were then imaged for TRITC-albumin fluorescence. All images shown include the tumor periphery. Bars = 100 μm. [Color figure can be viewed in the online issue, which is available at www.interscience.wiley.com.]

Download figure to PowerPoint

Tumor response to vascular-targeting PDT was monitored noninvasively by whole body fluorescence imaging. The EGFP-MatLyLu tumors were imaged for EGFP fluorescence before and after treatments. Representative tumor EGFP fluorescence images are shown in Figure 5. Control tumors grew rapidly and exhibited central necrosis when tumor reached about 8–10 mm in diameter. Dead EGFP-MatLyLu tumor cells were unable to produce EGFP, causing dead tumor tissues to appear as dark areas in the EGFP fluorescence images. PDT with 25 J/cm2 light dose induced a partial tumor necrosis, but this PDT condition failed to inhibit tumor growth (Fig. 5). In fact, tumor growth after this PDT treatment was even more rapid than control tumors and average tumor volume was nearly twice that of control tumors at 9 days after treatment (Fig. 6, p < 0.01). In contrast, the 50 J/cm2 PDT effectively inhibited prostate tumor growth as indicated by a substantial decrease in EGFP fluorescence (Fig. 5) and average tumor volume (Fig. 6, p < 0.01 compared to the control tumor) after treatment. EGFP fluorescence was barely detectable at 2 days after PDT. But small EGFP fluorescent spots, indicating the existence of viable tumor cells, were often found at tumor edges several days after treatment and gradually grew in size which led to tumor recurrence. As shown in Figure 7, some viable tumor cells were clearly detected in tumor periphery at 48 hr after 50 J/cm2 PDT.

thumbnail image

Figure 5. In vivo tumor EGFP fluorescence images showing tumor response to vascular-targeting PDT with verteporfin. The EGFP-MatLyLu tumors were treated with 25 or 50 J/cm2 light at 15 min after i.v. injection of 0.25 mg/kg dose of verteporfin. Tumor EGFP fluorescence was imaged daily for up to 9 days after treatment with a whole-body fluorescence imaging system as described in the Material and methods. Images at Day 0 were taken right before treatment. Control tumors received no treatment. Scale bar = 10 mm.

Download figure to PowerPoint

thumbnail image

Figure 6. Tumor volume changes after vascular-targeting PDT with verteporfin. The EGFP-MatLyLu tumors were treated with 25 or 50 J/cm2 light at 15 min after i.v. injection of 0.25 mg/kg dose of verteporfin. Control tumors received no treatment. Tumor volume at Day 0 represented the starting volume right before the treatment.

Download figure to PowerPoint

thumbnail image

Figure 7. H&E staining images showing the existence of viable tumor cells at the tumor periphery after vascular-targeting PDT with verteporfin. The EGFP-MatLyLu tumors were treated with 50 J/cm2 light at 15 min after i.v. injection of 0.25 mg/kg dose of verteporfin. H&E staining of tumor sections taken at 48 hr after treatment showed wide spread tumor cell death and vascular damage. But a small number of viable tumor cells were detected at the tumor periphery. Part of the image on the left, highlighted in the box, is shown at a higher magnification on the right. The letters V and D indicate the viable tumor area and the dead tumor area, respectively. Bars = 100 μm. [Color figure can be viewed in the online issue, which is available at www.interscience.wiley.com.]

Download figure to PowerPoint

Discussion

  1. Top of page
  2. Abstract
  3. Material and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

A whole-body animal fluorescence imaging system was used in this study to visualize noninvasively tumor response following PDT targeting of tumor blood vessels in an EGFP-expressing MatLyLu prostate tumor model. TRITC-albumin was used as a macromolecular probe to image tumor vascular barrier function (vascular permeability). The increase in the TRITC fluorescence intensity, caused by enhanced extravasation from blood vessels, is an indicator of vascular barrier disruption. Albumin has a plasma half-life of more than 24 hr and it was used to follow vascular permeability changes up to several hours after treatment.22

We found in the present study that vascular-targeting PDT increased vascular permeability in a dose-dependent manner, which is in agreement with our previous study and indicates that tumor vasculature is a primary target of PDT with verteporfin.6 Importantly, our results demonstrate that the enhanced TRITC-albumin tumor uptake as a result of PDT-induced permeability increase was not homogeneous in tumor tissues. Both in vivo and ex vivo tumor imaging studies indicate that increase in TRITC-albumin extravasation was significantly higher in the peripheral tumor area than in the interior tumor area. Because the accumulation of a circulating molecule in tumor tissues is dependent upon the existence of functional blood vessels, the enhancement of TRITC-albumin accumulation in the tumor periphery is likely related to the predominant localization of functional blood vessels in peripheral tumor areas after vascular-targeting PDT. As shown in Figure 4, PDT was remarkably effective in inducing interior tumor blood vessel shutdown while some peripheral vessels were still functional up to 24 hr after PDT. Early closure of central tumor vessels limited the enhancement of TRITC-albumin in the tumor interior, whereas prolonged perfusion of some peripheral tumor vessels allowed more TRITC-albumin to continuously extravasate in the tumor periphery. We and others have previously reported that peripheral tumor vessels tend to maintain perfusion function after vascular-targeting PDT.23–25 Our present results further demonstrate that continuous functioning of peripheral blood vessels, which had been permeabilized by PDT, led to preferential accumulation of circulating molecules in the tumor periphery.

The existence of functional blood vessels in the tumor periphery was associated with peripheral tumor cell survival after PDT. As shown in Figure 7, H&E staining indicated a rim of viable tumor cells in the tumor periphery at 48 hr after PDT in spite of extensive tumor necrosis. In vivo imaging of tumor EGFP fluorescence demonstrated that the survival of these peripheral tumor cells resulted in peripheral tumor recurrence (Fig. 5). Here we used EGFP as an indicator of tumor cell viability with the assumption that dead tumor cells are not able to synthesize EGFP and emit EGFP fluorescence. However, because EGFP has a half-life of more than 3 hr,26 monitoring EGFP fluorescence shortly after treatment might not accurately report tumor cell viability. Sufficient time is needed for the degradation of EGFP synthesized before treatment in order to use EGFP fluorescence to report cell viability. The observed decrease in EGFP fluorescence shortly after PDT in the present study was likely due to the oxidative degradation of EGFP during PDT rather than a real decrease in tumor cell viability. This was supported by the fact that there was little further decrease in EGFP fluorescence intensity over the following 5 hr period after PDT (Fig. 2c).

It is still not clear why peripheral tumor blood vessels react differently from interior blood vessels to vascular-targeting PDT. Understanding the mechanism behind this disparity in vascular response will help find ways to enhance the therapeutic effects of vascular-targeting PDT. Differences in vascular structure and function between tumor peripheral and interior blood vessels caused by morbid tumor pathobiology possibly contribute to such variations in vascular response. It is known that tumor tissues have higher tissue interstitial pressure than normal tissues because of leaky tumor blood vessels and poor lymphatic system function.27, 28 High tumor interstitial pressure is able to compress tumor vessels and lead to vessel collapse. Vessel compression and collapse are more severe in the tumor interior where tumor interstitial pressure is higher.29, 30 PDT has been shown to further increase tumor interstitial pressure as a result of enhancing vascular permeability.31, 32 Such an increase in tumor interstitial pressure will likely impose a greater compression on tumor blood vessels and cause vascular shutdown, especially in tumor interior areas. Moreover, we recently found that, compared to the interior tumor vessels, peripheral tumor blood vessels were generally larger and exhibited vascular lumen as well as more coverage of vascular pericytes and basement membrane.33 Less mechanic compression together with more vessel supporting structures might make peripheral tumor vessels more resistant than the interior vessels to vessel closure induced by vascular-targeting PDT.

The survival of peripheral tumor cells as a consequence of disparity in vascular response between peripheral and interior blood vessels represents a therapeutic challenge for the vascular-targeting PDT. Several strategies can be adopted to eliminate or at least minimize surviving tumor cells at the tumor periphery. First of all, we could increase the PDT dose to determine whether a higher dose of vascular-targeting PDT will lead to the shutdown of both interior and peripheral tumor blood vessels, resulting in an increased tumor cure. Secondly, as combination therapies have been routinely used in cancer treatments, one approach of enhancing photodynamic vascular targeting effectiveness is to combine it with other cancer therapies. Combination therapies can be designed based on different targeting principles. Targeting both tumor vascular and cellular compartments by combining vascular-targeting PDT with a cancer cell-targeted therapy could be a promising strategy because the increased vascular permeability induced by PDT has been shown to enhance drug delivery.6, 34, 35 Our present study further demonstrates that the enhancement of drug accumulation mainly occurred at the tumor periphery where tumor cell survival tends to occur after vascular-targeting PDT. Therefore, combining vascular-targeting PDT with other anticancer drug therapies will allow more anticancer agents to be preferentially deposited in the peripheral tumor area to kill tumor cells that otherwise might survive after PDT treatment.

In summary, we utilized in vivo animal fluorescence imaging combined with standard ex vivo tissue fluorescence microscopy to examine changes in vascular function and tumor cell viability after vascular-targeting PDT. Our results indicate that, although PDT causes an overall increase in vascular permeability, peripheral tumor blood vessels are somehow able to maintain perfusion function whereas interior blood vessels are shutdown shortly after PDT. Such a disparity in vascular response is conducive to peripheral tumor cell survival and also explains the preferential accumulation of circulating molecules in the tumor periphery. We are currently investigating the mechanisms underlying this response disparity and exploring therapeutic strategies that minimize the survival of peripheral tumor cells.

Acknowledgements

  1. Top of page
  2. Abstract
  3. Material and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References

The authors gratefully acknowledge Dr. Tayyaba Hasan of the Wellman Center for Photomedicine for helpful discussions and QLT Inc. for providing verteporfin.

References

  1. Top of page
  2. Abstract
  3. Material and methods
  4. Results
  5. Discussion
  6. Acknowledgements
  7. References
  • 1
    Dougherty TJ,Gomer CJ,Henderson BW,Jori G,Kessel D,Korbelik M,Moan J,Peng Q. Photodynamic therapy. J Natl Canc Inst 1998; 90: 889905.
  • 2
    Schmidt R. Photosensitized generation of singlet oxygen. Photochem Photobiol 2006; 82: 116177.
  • 3
    Chen B,Pogue BW,Hoopes PJ,Hasan T. Vascular and cellular targeting for photodynamic therapy. Crit Rev Eukaryot Gene Expr 2006; 16: 279305.
  • 4
    Brown SB,Mellish KJ. Verteporfin: a milestone in opthalmology and photodynamic therapy. Expert Opin Pharmacother 2001; 2: 35161.
  • 5
    Chen B,Pogue BW,Hoopes PJ,Hasan T. Combining vascular and cellular targeting regimens enhances the efficacy of photodynamic therapy. Int J Radiat Oncol Biol Phys 2005; 61: 121626.
  • 6
    Chen B,Pogue BW,Luna JM,Hardman RL,Hoopes PJ,Hasan T. Tumor vascular permeabilization by vascular-targeting photosensitization: effects, mechanism, and therapeutic implications. Clin Canc Res 2006; 12: 91723.
  • 7
    Liu DL,Svanberg K,Wang I,Andersson-Engels S,Svanberg S. Laser Doppler perfusion imaging: new technique for determination of perfusion and reperfusion of splanchnic organs and tumor tissue. Laser Surg Med 1997; 20: 4739.
  • 8
    Enejder AM,af Klinteberg C,Wang I,Andersson-Engels S,Bendsoe N,Svanberg S,Svanberg K. Blood perfusion studies on basal cell carcinomas in conjunction with photodynamic therapy and cryotherapy employing laser-Doppler perfusion imaging. Acta Derm Venereol 2000; 80: 1923.
  • 9
    Yu G,Durduran T,Zhou C,Wang HW,Putt ME,Saunders HM,Sehgal CM,Glatstein E,Yodh AG,Busch TM. Noninvasive monitoring of murine tumor blood flow during and after photodynamic therapy provides early assessment of therapeutic efficacy. Clin Canc Res 2005; 11: 354352.
  • 10
    Kruijt B,de Bruijn HS,van der Ploeg-van den Heuvel A,Sterenborg HJ,Robinson DJ. Laser speckle imaging of dynamic changes in flow during photodynamic therapy. Laser Med Sci 2006; 21: 20812.
  • 11
    Smith TK,Choi B,Ramirez-San-Juan JC,Nelson JS,Osann K,Kelly KM. Microvascular blood flow dynamics associated with photodynamic therapy, pulsed dye laser irradiation and combined regimens. Laser Surg Med 2006; 38: 5329.
  • 12
    Aalders MC,Triesscheijn M,Ruevekamp M,de Bruin M,Baas P,Faber DJ,Stewart FA. Doppler optical coherence tomography to monitor the effect of photodynamic therapy on tissue morphology and perfusion. J Biomed Opt 2006; 11: 044011.
  • 13
    Ohlerth S,Laluhova D,Buchholz J,Roos M,Walt H,Kaser-Hotz B. Changes in vascularity and blood volume as a result of photodynamic therapy can be assessed with power Doppler ultrasonography. Laser Surg Med 2006; 38: 22934.
  • 14
    Schmidt-Erfurth U,Niemeyer M,Geitzenauer W,Michels S. Time course and morphology of vascular effects associated with photodynamic therapy. Ophthalmology 2005; 112: 20619.
  • 15
    Zilberstein J,Schreiber S,Bloemers MC,Bendel P,Neeman M,Schechtman E,Kohen F,Scherz A,Salomon Y. Antivascular treatment of solid melanoma tumors with bacteriochlorophyll-serine-based photodynamic therapy. Photochem Photobiol 2001; 73: 25766.
  • 16
    Seshadri M,Spernyak JA,Mazurchuk R,Camacho SH,Oseroff AR,Cheney RT,Bellnier DA. Tumor vascular response to photodynamic therapy and the antivascular agent 5,6-dimethylxanthenone-4-acetic acid: implications for combination therapy. Clin Canc Res 2005; 11: 424150.
  • 17
    Yang M,Baranov E,Jiang P,Sun FX,Li XM,Li L,Hasegawa S,Bouvet M,Al-Tuwaijri M,Chishima T,Shimada H,Moossa AR, et al. Whole-body optical imaging of green fluorescent protein-expressing tumors and metastases. Proc Natl Acad Sci USA 2000; 97: 120611.
  • 18
    Hoffman RM. The multiple uses of fluorescent proteins to visualize cancer in vivo. Nat Rev Canc 2005; 5: 796806.
  • 19
    Ntziachristos V. Fluorescence molecular imaging. Annu Rev Biomed Eng 2006; 8: 133.
  • 20
    Nguyen TH,Oberholzer J,Birraux J,Majno P,Morel P,Trono D. Highly efficient lentiviral vector-mediated transduction of nondividing, fully reimplantable primary hepatocytes. Mol Ther 2002; 6: 199209.
  • 21
    Pogue BW,Gibbs SL,Chen B,Savellano M. Fluorescence imaging in vivo: raster scanned point-source imaging provides more accurate quantification than broad beam geometries. Technol Canc Res Treat 2004; 3: 1521.
  • 22
    Matsushita S,Chuang VT,Kanazawa M,Tanase S,Kawai K,Maruyama T,Suenaga A,Otagiri M. Recombinant human serum albumin dimer has high blood circulation activity and low vascular permeability in comparison with native human serum albumin. Pharm Res 2006; 23: 88291.
  • 23
    Chen B,Pogue BW,Goodwin IA,O'Hara JA,Wilmot CM,Hutchins JE,Hoopes PJ,Hasan T. Blood flow dynamics after photodynamic therapy with verteporfin in the RIF-1 tumor. Radiat Res 2003; 160: 4529.
  • 24
    Kurohane K,Tominaga A,Sato K,North JR,Namba Y,Oku N. Photodynamic therapy targeted to tumor-induced angiogenic vessels. Cancer Lett 2001; 167: 4956.
  • 25
    Koudinova NV,Pinthus JH,Brandis A,Brenner O,Bendel P,Ramon J,Eshhar Z,Scherz A,Salomon Y. Photodynamic therapy with Pd-bacteriopheophorbide (TOOKAD): successful in vivo treatment of human prostatic small cell carcinoma xenografts. Int J Canc 2003; 104: 7829.
  • 26
    Li X,Zhao X,Fang Y,Jiang X,Duong T,Fan C,Huang CC,Kain SR. Generation of destabilized green fluorescent protein as a transcription reporter. J Biol Chem 1998; 273: 349705.
  • 27
    Fukumura D,Jain RK. Tumor microenvironment abnormalities: causes, consequences, and strategies to normalize. J Cell Biochem 2007; 101: 93749.
  • 28
    Heldin CH,Rubin K,Pietras K,Ostman A. High interstitial fluid pressure - an obstacle in cancer therapy. Nat Rev Canc 2004; 4: 80613.
  • 29
    Rofstad EK,Tunheim SH,Mathiesen B,Graff BA,Halsor EF,Nilsen K,Galappathi K. Pulmonary and lymph node metastasis is associated with primary tumor interstitial fluid pressure in human melanoma xenografts. Canc Res 2002; 62: 6614.
  • 30
    Boucher Y,Jain RK. Microvascular pressure is the principal driving force for interstitial hypertension in solid tumors: implications for vascular collapse. Canc Res 1992; 52: 511014.
  • 31
    Fingar VH,Wieman TJ,Doak KW. Changes in tumor interstitial pressure induced by photodynamic therapy. Photochem Photobiol 1991; 53: 7638.
  • 32
    Leunig M,Goetz AE,Gamarra F,Zetterer G,Messmer K,Jain RK. Photodynamic therapy-induced alterations in interstitial fluid pressure, volume and water content of an amelanotic melanoma in the hamster. Br J Canc 1994; 69: 1013.
  • 33
    Chen B,He C,de Witte P,Hoopes PJ,Hasan T,Pogue BW. Vascular targeting in photodynamic therapy. In: HamblinMR,MrozP, eds. Advances in photodynamic therapy: basic, translational and clinicaled. Norwood, MA: Artech House Inc (in press).
  • 34
    Snyder JW,Greco WR,Bellnier DA,Vaughan L,Henderson BW. Photodynamic therapy: a means to enhanced drug delivery to tumors. Canc Res 2003; 63: 812631.
  • 35
    Debefve E,Pegaz B,Ballini JP,Konan YN,van den Bergh H. Combination therapy using aspirin-enhanced photodynamic selective drug delivery. Vascul Pharmacol 2007; 46: 17180.