Fibrinolytic Enzyme Cotherapy Improves Tumor Perfusion and Therapeutic Efficacy of Anticancer Nanomedicine

Poor intratumoral distribution of the drug carrier and inadequate drug delivery to tumors are significant challenges to successful translation of anticancer nanomedicine (1). Prolonging circulation times (2, 3) as well as optimizing particle size, shape, charge, and other physicochemical properties of the drug carrier may address these challenges (4), but only to a limited extent. The benefits of these approaches are limited by the erratic blood supply often observed in tumors.

Tumor perfusion is limited because of several peculiar characteristics of tumor blood vessels. These include inadequate coverage of the tumor, lack of transvascular pressure gradients, and their collapsed nature (5). Blood vessels in the core of the tumor are compressed under solid stress exerted by fast-dividing cells (6) and large amounts of extracellular matrix (ECM; ref. 7). Treatment with enzymes that degrade ECM components, such as collagen (8) and hyaluronic acid (9), has been shown to improve vascular characteristics and enhance blood supply in specific tumor types. However, the ubiquitous expression of collagen and hyaluronic acid in the body may limit the widespread use of these enzymes.

Because of the leaky nature of tumor blood vessels, fibrinogen, a soluble vascular protein, is deposited in the tumor matrix. The prothrombogenic activity of tumor cells leads to the conversion of fibrinogen to cross-linked fibrin, the principle ingredient of blood clot (10, 11). In fact, due to the constitutive activity of coagulation factors in tumors, they are often considered “overhealing wounds” (12). Presence of significant amounts of fibrin in the tumor matrix may contribute to increased solid stress often observed in tumors. In addition, the tumor-specific presence of fibrin makes it an attractive target for therapeutic interventions. However, the role of fibrin in limiting the intratumoral transport of drug carriers and the effect of fibrin degradation on their anticancer efficacy has not been previously studied.

We show here that administration of a fibrinolytic enzyme improves tumor perfusion by decompressing blood vessels and enhances chemotherapeutic activity of anticancer nanomedicine in syngeneic mouse melanoma and xenograft lung cancer models.

Tissue microarrays of various human tumor biopsies were obtained from Folio Biosciences. Anti-human fibrin(ogen) antibody was purchased from Dako. Poly(lactide-co-glycolide) (PLGA) was purchased from Lactel. l-lactide, bovine fibrinogen, and thrombin were obtained from Sigma Aldrich. Tissue plasminogen activator (tPA) was obtained from Abcam. Poly(ethylene glycol) was purchased from Laysan Bio Inc.

Tissue microarrays were digested with Proteinase K and stained with a polyclonal rabbit anti-human fibrin(ogen) antibody at a dilution of 1:1,000. Representative images were acquired using an Eclipse TS100 microscope (Nikon instruments) under ×400 magnification. Fraction of the area stained was quantified using ImageJ v1.48 software.

A block copolymer of poly(lactide) and poly(ethylene glycol) was synthesized via ring-opening polymerization reaction as described previously (13). The block copolymer was characterized using ¹H-NMR (Varian 400 MHz; ref. 13).

PLGA nanoparticles loaded with paclitaxel were synthesized by a single emulsion technique. Nanoparticles were surface functionalized with the block copolymer using the interfacial activity assisted surface functionalization technique developed in our laboratory. Nanoparticles were separated from free drug by repeated (three times) ultracentrifugation and resuspension in deionized water, and then lyophilized (13). PLGA nanoparticles loaded with a fluorescent dye (Coumarin 6) were synthesized similarly (14).

To determine particle size and zeta potential, nanoparticles were dispersed in deionized water and then subjected to dynamic light scattering analysis (Delsa Nano C, Beckman Coulter).

To determine drug loading, nanoparticles were dispersed in methanol (∼1 mg/mL) and the drug was extracted overnight. Nanoparticles were separated from extracted drug by centrifugation (14,000 rpm, 15 minutes). Drug concentration in the extract was analyzed using HPLC (Beckman Coulter; ref. 15).

A Transwell assay was used to determine the rate of migration of nanoparticles across fibrin matrices. In these studies, fibrin matrix was formed in situ in the top insert of a Transwell plate (12-mm diameter, 0.4-μm pore size, polycarbonate membrane). Fibrinogen (3 mg/mL) was dissolved in 0.9% w/v saline at 37°C. Plasminogen (0.1 U/mL) and thrombin (1 U/mL) were added to the fibrinogen solution. A 750-μL aliquot of the solution was then quickly transferred to the inserts. The plate was placed in a cell culture incubator overnight to allow fibrin gel formation.

At the beginning of the experiment, the bottom well was filled with 1 mL of 0.15 mmol/L PBS (pH 7.4, 1× PBS). Nanoparticles (dispersed in deionized water at a concentration of 5 mg/mL; 0.1 mL) were mixed with different amounts of tPA and added on top of the fibrin matrices. At various time points, contents of the bottom well were collected and lyophilized. The bottom well was replenished with fresh 1× PBS. The lyophilized samples were extracted overnight with methanol. Concentration of paclitaxel in the extract was determined using HPLC.

To determine the lag time for the appearance of drug in the bottom chamber, a curve given by the following equation was used to fit the data:

where A is the cumulative amount of drug in the bottom chamber, |{A_0}$| is the amount of drug added to the top chamber, k is the rate constant for transport across the insert, and |{t_{lag}}$|is the lag time. H(t) is a Heaviside step function described as:

Model fitting was performed using Matlab software (Mathworks).

We also calculated the apparent permeability (Papp) of drug into the bottom chamber. Apparent permeability was calculated as described by Hubatsch and colleagues (16) using the formula

where dQ/dt is the mass of paclitaxel transported per unit time after the lag time, A is the surface area of the insert, and C₀ is the initial concentration of paclitaxel in the top insert.

B16F10 murine melanoma cells were provided by Prof. Olin (University of Minnesota, Minneapolis, MN). Olin's laboratory obtained the cells from the ATCC with authentication by short tandem repeat. A549 human lung adenocarcinoma cells transfected with firefly luciferase gene (A549-luc) were purchased from Caliper Life Sciences. Both cells were cultured in RPMI1640 medium supplemented with 10% v/v FBS and 1% v/v penicillin and streptomycin under humidified atmosphere with 5% CO₂ at 37°C.

Cellular uptake of nanoparticles across fibrin matrices was determined using a Transwell assay. Fibrin matrices were formed in the top inserts of the Transwell plate as described above. B16F10 cells were seeded in the bottom well (1 × 10⁶ cells/well) and allowed to adhere overnight. On the day of the experiment, media were replaced with serum-free media (1 mL/well). Coumarin 6–loaded nanoparticles (0.25 mg dispersed in 0.05 mL deionized water) were mixed with increasing amounts of tPA and added on top of the fibrin matrix. After 4 hours, cells were washed with cold 1× PBS, and trypsinized. The cells were then transferred to flow tubes and centrifuged at 1,000 rpm for 5 minutes. The supernatant was removed and cells were suspended in cold 1× PBS. Cellular fluorescence was analyzed using flow cytometry (BD FACSCalibur flow cytometer).

All animal studies were performed in compliance with protocol approved by the Institutional Animal Care and Use Committee at the University of Minnesota (Minneapolis, MN). The efficacy of tPA–paclitaxel combination therapy was evaluated in a mouse xenograft model of human lung cancer. The lung tumor model was set up as described previously (17), with some minor changes. A549-luc cells (∼6 × 10⁵ cells/mouse) were dispersed in 1× PBS and injected in 4- to 6-week-old, female SCID mice via the lateral tail vein. Tumor growth in the lungs was determined by monitoring the bioluminescence using IVIS spectrum in vivo imaging system.

About 4–5 weeks after cell injection, animals were treated intravenously with saline or paclitaxel (40 mg/kg) or a combination of paclitaxel and tPA (436 μg/kg). The treatments were administered thrice every 96 hours. Tumor bioluminescence was monitored every 3–4 days.

B16F10 cells (1 × 10⁶ cells dispersed in 0.05 mL 1× PBS) were injected subcutaneously in 4- to 6-week-old female C57BL/6 mice. Tumor dimensions were measured daily using a digital caliper. Tumor volume was calculated as 0.5 × length × width². Treatment was initiated when the tumor volume reached approximately 100 mm³. Animals were treated intravenously with saline or tPA (436 μg/kg) or paclitaxel nanoparticles (40 mg paclitaxel/kg) or a combination of paclitaxel nanoparticles and tPA. Animals were treated with three doses administered 96 hours apart.

Tumors from the B16F10 efficacy study were collected and fixed in 5% formalin in PBS for 48 hours. The tumors were then removed from the formalin solution, washed thrice with 1× PBS, and stored in 70% ethanol until further processing. Tumor sections were stained with a goat anti-mouse CD31 antibody (Santa Cruz Biotechnology).

Fifty random images of each tumor section were obtained using an optical microscope at ×400 magnification. Diameters of the CD31⁺ blood vessels were measured using ProgRes Capture software (Jenoptik AG). On an average, each image contained 2–3 blood vessels. Six sections from each treatment group (2 sections × 3 animals/group) were analyzed. Hence, the results represent approximately 600–900 blood vessels/treatment group.

C57BL/6 mice were shaved and inoculated with B16F10 cells as described in the efficacy study. When tumors reached a volume of approximately 100 mm³, animals were randomly assigned to two groups and treated with three doses of either saline or tPA (436 μg/kg) every 96 hours. About 1 hour after each treatment, tumor perfusion was analyzed using ultrasound imaging.

Ultrasound imaging was performed using a Vevo 2100 system (FujiFilm VisualSonics). Animals were anesthetized using isoflurane, and were maintained on the gas throughout the imaging. Any hair on the tumor was removed using Nair (Church and Dwight). Using a motorized stage, an MS550 probe was placed on top of the tumor. An appropriate field of view was determined by monitoring tumor boundary and location in B-mode. Perfusion area was visualized by switching to the Power Doppler mode. Power Doppler images were analyzed using the ImageJ v1.48 software.

C57BL/6 mice bearing B16F10 tumors were used for these studies. Tumors were established as described before. When the tumor volume reached approximately 100 mm³, animals were treated with either saline or tPA (436 μg/kg). Four days later, animals were treated with Doxil (20 mg doxorubicin/kg) or a combination of Doxil and tPA. On the next day, animals were sacrificed and their tumors were collected. Tumors were embedded in Tissue-Tek optimum cutting temperature (OCT) compound (Sakura Finetek USA, Inc.) and flash frozen in a slurry of dry ice in isopentane. Liver, lungs, spleen, and brain were harvested and processed similarly.

Tumor or tissue sections (5-μm thickness) were cut and analyzed using a Nikon A1Rsi confocal super resolution microscope (Nikon Instruments Inc.). Fluorescence and transmitted light images were obtained at ×10 magnification. For fluorescence imaging, samples were excited using a 488-nm laser, and emitted light was collected at 595/50 nm. Using a motorized stage, serial images were obtained across the entire tumor section. Images were stitched together using NIS Elements Viewer 4.0 software (Nikon Instruments Inc.). A region of interest (ROI) was drawn manually around the periphery of the tumor, and total fluorescence intensity and area of the ROI was measured. The ROI was shrunk to approximately 40% and 10% of its original area, and similar measurements of fluorescence intensity and area were carried out. Fluorescence intensity normalized to area in the central 40% and central 10% of the tumor section relative to total fluorescence intensity was measured and used as an indicator of penetration of the drug into the core of the tumor.

Statistical analyses for matrix migration and cell uptake studies were performed using one-way ANOVA. For efficacy studies and ultrasound analysis, statistical significance of the differences between two treatment groups was determined using the Student t test. For all studies, P < 0.05 was considered statistically significant.

All other methods are described in Supplementary Information.

Several solid tumors have leaky blood vessels. This leads to the deposition of vascular proteins in the tumor matrix. We were particularly interested in analyzing the deposition of fibrin, an important vascular protein involved in blood clotting.

We examined the occurrence of fibrin in an array of human tumor biopsies. We found that all the tumors considered in our studies showed significant fibrin deposition (Fig. 1A–H). Fraction of the tumor area stained for fibrin ranged from approximately 20% to 90% (Fig. 1I). We next determined whether fibrin levels were affected by the stage of the disease, metastatic status, or the type of cancer. To minimize variability, we considered only lung carcinomas. We found that both adenocarcinomas and squamous cell carcinomas had comparable amounts of fibrin (Fig. 1J). These levels were unaffected by the stage of the disease (Fig. 1K) or the presence of lymph node metastasis (Fig. 1L). Representative images of lung carcinomas in different stages of progression are shown in Supplementary Fig. S1.

We also analyzed the expression of fibrin in several human and murine tumors grown in mice. Similar to our results with the human tumor samples, high levels of fibrin were found in these tumors as well (Supplementary Fig. S2). These results corroborated well with previous findings that fibrin is an important component of the tumor ECM (18, 19).

Previous reports have shown that degrading other components of tumor ECM, such as hyaluronic acid and collagen, can improve delivery of drugs to tumor cells. This result is manifested as both enhanced intratumoral distribution of the drug as well as improved tumor perfusion (9, 20, 21). We determined whether degrading fibrin in tumors had a similar effect. To degrade fibrin within tumors, we used tPA, an enzyme that is used clinically to treat patients with pulmonary embolism, myocardial infarction, and stroke (22–24).

PLGA nanoparticles loaded with paclitaxel and surface-functionalized with poly(ethylene glycol) were used as model drug carriers. The results of the physicochemical characterization of the nanoparticles are summarized in Supplementary Information (Supplementary Table S1).

To study the diffusion of nanoparticles in fibrin matrices, we used a Transwell assay. We found that the mobility of nanoparticles was significantly compromised by the matrix. No drug was detected in the receiver chamber through the time course of the experiment (Fig. 2A). However, addition of tPA significantly improved the movement of nanoparticles. With an increase in the amount of tPA, the lag time (t_lag) for the appearance of drug in the receiver chamber decreased significantly (Fig. 2B). To determine the rate of drug transport from the top to bottom chamber, we calculated Papp (Fig. 2C). In the absence of tPA, there was no transport and Papp was zero. In the presence of 0.05 μg of tPA, Papp of the drug was 0.01 cm/h. With an increase in the amount of tPA to 0.5 μg, Papp nearly doubled. However, further increase in tPA led to no change in Papp.

We also analyzed the cellular uptake of nanoparticles across fibrin gels. In these studies, nanoparticles were separated from a monolayer of B16F10 cells by fibrin matrix. As a positive control, we added nanoparticles directly to the cells. This resulted in a rapid uptake of nanoparticles. However, in the presence of the fibrin matrix, nanoparticle uptake was significantly reduced (Fig. 2D). Addition of tPA resulted in a dose-dependent increase in cell uptake (Fig. 2D and E). tPA had no effect on cellular uptake of nanoparticles in the absence of the fibrin gel (Supplementary Fig. S3).

These experiments show that fibrin matrix could significantly retard the movement of nanoparticles, and this effect could be reversed, at least partially, by cotreatment with tPA.

We determined the safety and tolerability of a combination of paclitaxel nanoparticles and tPA. Treatment with paclitaxel (as single therapy or in combination) led to a slight loss in body weight (∼5%), which was recovered within 48 hours of dosing (Supplementary Fig. S4A and S4B). The levels of all the markers of liver function were elevated in response to paclitaxel therapy. Coadministration of tPA did not have a significant impact on the body weight or the level of these markers (Supplementary Fig. S4A–S4D)

We further evaluated the safety of tPA, and its effect on fibrin. Lung tumor–bearing mice were treated with tPA, and blood D-dimer (a degradation product of fibrin) levels were analyzed using ELISA. In tumor-bearing mice, treatment with tPA led to an increase in plasma levels of D-dimer protein. The increase in D-dimer levels was observed in tumor-bearing mice but not in healthy mice (Supplementary Fig. S5). This suggests that tPA specifically degrades fibrin in tumor-bearing animals and does not have any detectable thrombolytic effect in healthy animals, providing further evidence to its safety.

We tested the efficacy of the combination of tPA and paclitaxel nanoparticles in a mouse orthotopic model of human lung cancer. Animals treated with paclitaxel in solution had a slower tumor growth rate as compared with saline-treated animals. Paclitaxel in nanoparticles reduced the tumor growth rate even further. However, greatest tumor growth inhibition was seen when paclitaxel nanoparticles were administered in combination with tPA (Fig. 3A).

We further tested the effect of tPA on the anticancer activity of paclitaxel in a syngeneic mouse melanoma model. Treatment with tPA or paclitaxel nanoparticles reduced tumor growth rate as compared with saline controls. Importantly, combining paclitaxel nanoparticles with tPA led to a further improvement in chemotherapeutic efficacy (Fig. 3B). However, the improvement in efficacy with tPA coadministration was observed only at early time points in this more aggressive model.

Blood vessels in many solid tumors are compressed because of the presence of copious amounts of ECM components and a fast-growing cell population. Previous experiments have shown that the administration of cytotoxic therapies (6, 25) or ECM-degrading enzymes (9) can decrease solid stress and restore vascular function. We hypothesized that degrading fibrin will have a similar effect.

Similar to that reported for other solid tumors (9), B16F10 tumors also had a significant number of collapsed blood vessels (Fig. 4A and D). Most blood vessels were <10 μm in diameter. After treatment with paclitaxel nanoparticles, there was an increase in the median blood vessel diameter (Fig. 4C and E). When treated with a combination of paclitaxel nanoparticles and tPA, the median vessel diameter increased even further (Fig. 4B, C, and F). It is interesting to note that regardless of the treatment, the diameter of most blood vessels was <10 μm. However, treatment with paclitaxel alone or in combination with tPA led to an increase in the number of blood vessels in the 10- to 20-μm diameter range (Fig. 4D–F).

We further obtained real-time physiologic information regarding blood perfusion in response to fibrinolytic therapy. Ultrasound imaging was used to track blood vessels in the subcutaneous B16F10 tumors. In saline-treated animals, tumor perfusion area reached a maximum of approximately 1% after the second dose (Fig. 4G and I). At all time points analyzed, tPA-treated tumors displayed a greater perfusion area (∼3- to 4-fold higher) as compared with the saline-treated tumors. In animals treated with tPA, tumor perfusion area was maximal after the second dose (∼4% of tumor area; P < 0.05 vs. saline-treated; ref. Fig. 4H and I).

We determined the effect of tPA administration on the intratumoral distribution of nanocarriers. Doxil was used as a model anticancer nanomedicine in these studies because fluorescence of doxorubicin allows visualization of the drug in tissue sections.

In tumors treated with Doxil, fluorescence intensity was higher toward the periphery and minimal in the interior regions of the tumor (Fig. 5A). These results are in agreement with recent studies that show high drug accumulation in the tumor periphery as compared with that in the central core of the tumor (26). In tumors treated with a combination of tPA and Doxil, fluorescence intensity was still maximal toward the periphery. However, compared with the control Doxil group, greater distribution of doxorubicin-associated fluorescence intensity could be observed in the tumor interior (Fig. 5B and C). These results indicate that treatment with tPA resulted in improved intratumoral distribution of Doxil.

Interestingly, the distribution of Doxil-associated fluorescence intensities in liver, lungs, spleen, and brain cross sections were homogenous (Fig. 5D–G; Supplementary Fig. S6). This is in contrast with tumor sections that showed significantly lower fluorescence intensity in the core. Treatment with tPA did not affect the intratissue distribution of fluorescence. The total fluorescence intensity in tumors and other tissues was comparable between the Doxil- and Doxil + tPA–treated groups (Supplementary Fig. S7). This suggests that the overall biodistribution of Doxil is not affected by tPA cotherapy.

Solid tumors are characterized by nonhomogenous blood supply and the presence of varying degrees of hypoxia. These hypoxic regions often harbor aggressive and drug-resistant tumor cells (27). Thus, achieving therapeutic concentrations of the drug in these under-supplied regions of the tumor is a significant challenge.

Tumor blood vessels are formed in response to proangiogenic factors secreted by tumor cells (28). An imbalance in pro- and antiangiogenic factors within the tumor tissue results in the formation of blood vessels that are abnormal and porous (29). Fluid leakage from these porous blood vessels and lack of functional lymphatics in the core of the tumor result in fluid accumulation in the tumor (30). This leads to diminished transvascular pressure gradient in certain regions of the tumor (30). In addition, because of the presence of fast-dividing cells within a limited volume and a highly dense ECM, blood vessels in the core of the tumor are compressed and not fully functional (7). Consequently, drug exchange through convection is highly compromised (5). Improving fluid dynamics in the tumor is, therefore, likely to have a significant impact on the delivery of drugs to tumors.

Several strategies have been used for improving drug transport within the tumor (31). The use of antiangiogenics results in the normalization of blood vessels, leading to reduced fluid leakage out of the blood vessels, and a decrease in interstitial fluid pressure (29). Consequently, convectional transport across blood vessels increases. On the other hand, decreasing the production or degrading the ECM in the tumor has also been shown to improve drug delivery. Decreasing the density of ECM can improve the diffusivity of the carrier/drug in the tumor matrix. Moreover, this can lead to decompression of the collapsed blood vessels, resulting in increased tumor perfusion and improved drug delivery (8, 9).

Because of the leaky nature of tumor blood vessels, there is significant fibrinogen deposition within the tumor. The prothrombogenic activity of the tumor microenvironment leads to the activation of fibrinogen to fibrin. The role of fibrin in tumor growth and metastasis has been well studied (32–35). During the initial stages of tumor growth, fibrin provides a scaffold for the growth of tumor cells. Fibrin also plays an important role in angiogenesis, modulating the influx of macrophages and in the storage of growth factors (36, 37). Depletion of fibrinogen may reduce formation of pulmonary metastatic foci. This effect is likely due to the loss of fibrinogen-mediated protection of tumor cells against natural killer cells (34).

However, the role of fibrin in mediating solid stress and limiting drug delivery to tumors has not been studied extensively. In this study, we used enzymatic degradation of fibrin to improve the delivery and chemotherapeutic efficacy of paclitaxel. Through IHC studies, we found that treatment with tPA led to an increased blood vessel diameter. These data were supported by increased tumor perfusion as noted by ultrasound imaging. Importantly, coadministration of tPA led to an improved chemotherapeutic activity of paclitaxel nanoparticles in mouse models of melanoma and orthotopic lung cancer. A recent study analyzed the effect of tPA on the anticancer effect of nanoparticles in a xenograft subcutaneous A549 model (38). In agreement with our results, treatment with tPA led to an improved chemotherapeutic activity of nanoparticles. The benefit of cotreatment with tPA was different in the two tumor models used in our study. Thus, the type and location of the tumor may play a critical role in determining the improvement in activity obtained with tPA. Studying these effects in a transgenic tumor model or patient-derived tumor models will likely provide additional mechanistic insights.

Previous studies have analyzed the expression of fibrin in animal tumor models (18, 19). However, a systematic analysis of fibrin expression in human tumors has not been performed previously. Fibrin levels in human tumor biopsies were variable, with no tumor type showing either a complete lack or consistently high level of fibrin. There are several reasons cited for this heterogeneity in fibrin expression. Current IHC techniques cannot distinguish between fibrinogen and cross-linked fibrin. In addition, bleeding during the collection of the tumor biopsy can also significantly affect the results (36). Accurately quantifying fibrin levels in tumors is important to successfully advance adjuvant fibrinolytic therapy. Novel peptides that specifically identify fibrin could be useful in this regard (39, 40).

We used a Transwell assay to measure the mobility of nanoparticles through fibrin gels, and the influence of tPA on this mobility. Transwell assays have been employed previously to characterize nanoparticle/drug transport across cell monolayers. In those assays, nanoparticle/drug transport commences upon their introduction into the top chamber. In contrast, in our experiment, due to the presence of a dense matrix of fibrin, drug was not detected in the bottom chamber over the course of the study (10 hours). Addition of tPA reduced the time to appearance of drug in the bottom chamber in a dose-dependent manner. This indicates that fibrin gels used in our studies are resistant to nanoparticle transport. In addition, transport across fibrin gels occurred in two phases. In the first phase characterized by digestion of fibrin gels by tPA, no drug deposition was seen in the bottom chamber. In the second phase, once the fibrin was digested, drug transport occurred at a constant rate. To quantify transport in both these phases, we used two parameters viz. lag time and apparent permeability. Lag time represents the time it takes for the gel to be digested, and apparent permeability represents the drug transport after fibrin digestion.

The clinical use of anticoagulants and fibrinolytic therapies in patients with cancer has been reported before (41–45). These studies have shown that the administration of anticoagulants, such as heparin and warfarin, improve mean survival in patients with small-cell lung cancer. The rationale presented for the use of anticoagulants is that it reduces the occurrence of spontaneous thromboembolism and tumor cell dissemination (44, 46). Some studies have reported the beneficial effects of using urokinase in conjunction with chemotherapy (41). However, the mechanism for the improved response to chemotherapy was not explored. In our studies, the benefit of tPA on the anticancer drug activity manifested itself through increased tumor vascularity, improved perfusion, and more uniform distribution of the drug within the tumor.

The use of tPA is contraindicated in patients with acute intracranial hemorrhage, severe uncontrolled hypertension, head trauma or stroke in previous three months, thrombocytopenia and coagulopathy, concomitant anticoagulant therapy, severe hypoglycemia, or hyperglycemia (47). Patients with these coexisting pathologies should be excluded from tPA cotherapy. Also, the presence of intracranial neoplasm is a contraindication for tPA therapy (48). However, tPA is not contraindicated for other cancers. Interestingly, there are several reports of patients with intracranial neoplasms who received intravenous thrombolysis but did not have intracranial hemorrhage (47, 49–51). On the basis of these reports, a recent review article (47) concluded that intravenous thrombolysis can be safe in patients with intracranial neoplasms, provided the lesion is small and extra-axial in location. However, a careful assessment of benefit-to-risk ratio and comorbidities is imperative before the clinical adoption of this strategy.

The time course of change in perfusion area with tumor volume is interesting. Perfusion area in untreated tumors and tPA-treated tumors was highest after the second dose of tPA. Previous work has shown that blood flow generally decreases with an increase in tumor volume (52). The disparity in results may be due to different techniques used for measurement of blood flow. Lack of sensitivity of the ultrasound technique may have limited our ability to detect blood vessels in small tumors.

Studies with antiangiogenic therapy have shown that there is a time window associated with the normalization of blood vessels (53, 54). In other words, the decrease in interstitial fluid pressure is transient. The improvement in drug delivery is maximal in this time window. It is conceivable that a similar phenomenon may occur with tPA treatment as well. Modeling reports suggest that an increased perfusion in leaky blood vessels can result in a rapid increase in interstitial fluid pressure (55). This would quickly neutralize the advantage obtained with increased perfusion. Moreover, tPA has a short half-life. If the deposition of fibrin in tumors is rapid, the decrease in fibrin content of the tumor will likely be observed only for a short period of time. The pharmacokinetics of nanoparticles and tPA accumulation in the tumor may also be different. Additional studies are needed to determine the optimal frequency of tPA dosing and the time window following tPA dosing when the improvement in drug delivery is maximal. Furthermore, we have only considered PLGA nanoparticles in this study. The effect of tPA codelivery on other nanoparticulate systems of varying sizes would be of interest as well. A careful consideration of all these aspects can help maximize therapeutic benefits obtained from this strategy.

Poor vascular architecture and the consequent inadequate drug delivery limit the overall efficacy of chemotherapy. Fibrin deposition within the tumor can be an important cause for the compression of tumor blood vessels. Administration of tPA improved blood supply in solid tumors and enhanced the chemotherapeutic activity of paclitaxel-loaded polymeric nanoparticles.

A.R. Kirtane, T. Sadhukha, and J. Panyam have ownership interest (including patents) in use of fibrinolytic enzymes to improve chemotherapeutic activity of nanoparticles. T. Sadhukha is a senior research scientist in Albany Molecular Research Inc. No potential conflicts of interest were disclosed by the other authors.

Conception and design: A.R. Kirtane, T. Sadhukha, J. Panyam

Development of methodology: A.R. Kirtane, T. Sadhukha, J. Panyam

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): A.R. Kirtane, V. Khanna, B. Koniar

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): A.R. Kirtane, H. Kim, V. Khanna, J. Panyam

Writing, review, and/or revision of the manuscript: A.R. Kirtane, J. Panyam

Study supervision: J. Panyam

We thank Drs. Sveotmir Markovic (Mayo Clinic) and Vladimir R. Muzykantov (University of Pennsylvania, Philadelphia, PA) for helpful discussions on the clinical use of fibrinolytics. The authors would like to thank Josh Parker (Comparative Pathology Shared Resource) for IHC analysis. Authors also acknowledge the Minnesota Supercomputing Institute for providing access to Matlab software. Live animal imaging (bioluminescence and ultrasound imaging) and confocal microscopy was performed at the University Imaging Center at the University of Minnesota (Minneapolis, MN). The authors also thank Mayank Verma and Dr. Guillermo Marques for providing training on ultrasound imaging and confocal microscopy.

This work was funded by the Center for Pharmaceutical Development, University of Minnesota (J. Panyam), and the Doctoral Dissertation Fellowship, University of Minnesota (A.R. Kirtane).

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.