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The therapeutic utility of cytochrome P450-based enzyme prodrug therapy is well established by preclinical studies and in initial clinical trials. The underlying premise of this gene therapy is that intratumoral P450 expression leads to in situ activation of anti-cancer P450 prodrugs, such as cyclophosphamide (CPA), with intratumoral accumulation of its activated, 4-OH metabolite. In mice bearing 9L gliosarcomas expressing the CPA 4-hydroxylase P450 2B6, enhanced tumor apoptosis was observed 48h post-CPA treatment, however, intratumoral 4-OH-CPA levels were indistinguishable from those of P450-deficient tumors, indicating that the bulk of activated CPA is derived from hepatic metabolism. In contrast, in 9L tumors expressing P450 2B11, a low Km CPA 4-hydroxylase, intratumoral 4-OH-CPA levels were higher than in blood, liver, and P450-deficient tumors. Intratumoral 4-OH-CPA increased dose-dependently, without saturation at 140mg/kg CPA, suggesting restricted tumor cell permeation of the parent drug. To circumvent this problem, CPA was administered by direct intratumoral injection, which increased the Cmax and AUC of intratumoral 4-OH-CPA by 1.8- and 2.7-fold, respectively. An overall 3.9-fold increase in intratumoral 4-OH-CPA AUC and increased anti-tumor activity were obtained when CPA release to systemic circulation was delayed using the slow-release polymer poloxamer 407 (F127) as vehicle for intratumoral CPA delivery. These findings highlight the advantage of gene therapy strategies that combine low Km P450 prodrug activation enzymes with slow, localized release of P450 prodrug substrates.
The chemosensitization of tumor cells by intratumoral activation of anticancer prodrugs is an emerging adjuvant therapy that can increase the efficacy of several commonly used anti-cancer agents. Intratumoral prodrug activation may be enhanced by introduction of prodrug-activation enzymes of bacterial, viral, or mammalian origin using a variety of gene therapy vectors, including retoviruses, adenoviruses, herpes viruses, as well as non-viral vectors 1–3. Chemosensitization is achieved when an anticancer prodrug is activated locally in tumor cells, which augments cytotoxic responses, both in the prodrug-activating tumor cell and in naive bystander tumor cells exposed to diffusible cytotoxic metabolites.
One such prodrug activation therapy utilizes genes that encode mammalian cytochrome P450 enzymes 4, 5. Drug metabolizing P450 enzymes are highly expressed in the liver and are generally found at very low levels in human tumors 6, 7. These enzymes catalyze the metabolism of more than 35 anticancer drugs 8, 9, of which 12 drugs, including the widely used alkylating agent prodrug cyclophosphamide (CPA) and its isomer ifosfamide (IFA), are activated via P450 metabolism 10. CPA is used in both adjuvant and high-dose chemotherapy settings and is effective against a broad range of tumors, including breast cancer and lymphomas 11. Two hepatic cytochromes P450, P450 2B6 and P450 3A4, catalyze a substantial fraction of CPA activation in human liver with electron input from NADPH P450 reductase 12. CPA is converted to 4-OH-CPA, a cell cycle-independent alkylating metabolite that readily diffuses out of hepatocytes and can exert cytotoxicity at distant sites, including tumor and sensitive host tissues. This intrinsic diffusibility of 4-OH-CPA underlies the bystander cytotoxic response that is obtained when CPA is combined with P450-prodrug gene therapy 13. The efficacy of P450 gene therapy can be enhanced in several ways, including combination with bioreductive prodrugs that are activated by P450 and/or P450 reductase 14, 15, by using a metronomic, anti-angiogenic CPA treatment schedule 16, 17 and by using tumor cell-replicating herpes virus 18 and adenovirus 19 to spread the therapeutic P450 gene and augment tumor cell lysis.
In the present study we investigated the activation of CPA in vivo in solid tumors that express P450 in an effort to improve the pharmacokinetics of CPA metabolism and enhance the efficacy of this P450 prodrug-activation strategy. Our findings demonstrate that despite an increase in chemosensitivity, tumor cell expression of the high Km CPA 4-hydroxylase enzymes P450 2B6 and P450 2B1, used in recent clinical trials 20, 21, does not lead to a measurable increase in intratumoral 4-OH-CPA levels. In contrast, introduction of P450 2B11, a low Km CPA 4-hydroxylase 22, increases the intratumoral concentration of 4-OH-CPA above that found in P450-deficient tumors. Further improvements in intratumoral pharmacokinetics and increased net tumor cell exposure to 4-OH-CPA were obtained by direct intratumoral CPA delivery via a slow-release polymer, substantially enhancing the overall anti-tumor response.
CPA, chloroquine and puromycin were purchased from Sigma-Aldrich (St. Louis, MO). 4-hydroperoxy-CPA was obtained from ASTA Pharma (Bielefield, Germany). IFA was obtained from the Drug Synthesis and Chemistry Branch of the National Cancer Institute (Bethesda, MD). Fetal bovine serum (FBS) and DMEM were purchased from Life Technologies (Grand Island, NY). F127, a 12,600 average molecular weight ethylene oxide and propylene oxide block copolymer 23 (also known as Pluronic F127, poloxamer 407 and Pluracare F-127 Prill Surfactant), was obtained from BASF Corporation (Florham Park, NJ) and contained 100 ppm BHT. Rat 9L gliosarcoma cells were grown in a humidified, 5% CO2 atmosphere at 37°C in DMEM containing 10% FBS, 100 units/mL penicillin, and 100 µg/mL streptomycin. Casputin (Cat. SE760) and the caspase 3 substrate Ac-DEVD-AMC (Cat. P-411) were purchased from Biomol International LP (Plymouth Meeting, PA).
9L gliosarcoma cells were infected with pBabe-based retrovirus expressing P450 2B1, P450 2B6 or P450 2B11 cDNA linked to P450 reductase cDNA via an internal ribosome entry site sequence (9L/2B1, 9L/2B6 and 9L/2B11 cells, respectively) as described 24. 9L cells infected with an empty pBabe-based retrovirus were used as P450-deficient controls. Immunochemical analysis using anti-P450 2B antibodies revealed P450 2B expression in ~90–95% of the cultured 9L/2B cells. This percentage decreased to ~50% or less of the cells when grown s.c. as solid tumors (J Ma and DJ Waxman, unpublished observations). 9L tumor cells were grown to near-confluence in DMEM culture medium containing 10% FBS, harvested by trypsin digestion, washed once and resuspended in FBS-free DMEM and placed on ice. Five week-old (23–25 g) male ICR/Fox Chase outbred immunodeficient scid mice (Taconic Farms, Germantown, NY) were injected on each flank with 4 × 106 9L, 9L/2B1, 9L/2B6, or 9L/2B11 cells in 0.2 ml of FBS-free DMEM using an 0.3 ml U-100 insulin syringe and a 28G-1/2 needle. All mice were housed and treated in the Boston University Laboratory of Animal Care Facility using approved animal protocols.
Pharmacokinetic data were collected when the tumors reached 800–1000 mm3 in size. Mice were given a single injection of CPA or IFA at doses specified in each experiment, either by systemic (i.p.) injection or by direct intratumoral (i.t.) injection as described below. In some cases, CPA was dissolved in 0.2% NaCl or in a 23% solution of F127 dissolved in 0.2% NaCl and then injected i.t. as specified in each experiment. The final CPA concentration was adjusted to 12.5–14 mg/ml based on each individual mouse’s body weight to keep the injected volume (60 µl per tumor) and the CPA dose (50 mg/kg BW) constant. Intratumoral CPA delivery was achieved using a syringe pump (model KDS100, KD Scientific, Holliston, MA) set a 1 µl/sec with a 30-gauge needle (20 µl per injection × 3 injection sites per tumor). A 23% F127 polymer solution was prepared as follows: 3 g of F127 was dissolved in 7.0 ml of sterile 0.2% NaCl with shaking at 4°C over a 24 h period to obtain a 30% F127 solution. A sterile solution of 60 mg/ml CPA was sonicated before use and then mixed with the required vol of 30% F127 stock solution to achieve a final concentration of 23% F127. Thus, for a 30 g mouse to be treated with 1.5 mg CPA (i.e., 50 mg CPA/kg BW), 208 µl of 60 mg/ml of CPA solution was mixed with 767 µl of 30% F127 and 25 µl of 0.2% NaCl. A vol of 60 µl was then injected directly into each tumor. Mice were killed 6, 15, 30, 60, 120, and 240 min after CPA treatment (n = 3 mice/time point).
4-OH-CPA levels were assayed in 200–400 µl of blood collected by heart puncture using a syringe containing 10 µl of heparin at 100 U/ml and 2–4 µl of 0.5 M semicarbazide (final concentration 5 mM) to stabilize the 4-OH-CPA. The concentration of 4-OH-CPA in liver and in tumor was determined using ~ 0.4 g of tissue pre-washed with 0.1 M KPi buffer, pH 7.4 containing 0.1 mM EDTA and 5 mM semicarbazide. Tissues were homogenized using a Polytron homogenizer, model PT-2000 (Kinematica, Luzern, Switzerland) in 0.1 M KPi buffer containing 0.1 mM EDTA and 5 mM semicarbazide (5 ml per g tissue). Homogenized blood, liver and tumor samples were centrifuged in Sorvall SA-600 rotor at 3000 rpm for 30 min at 4°C, and supernatants were stored at −80°C prior to HPLC analysis.
Extracts were prepared from 9L and 9L/2B6 tumors (880 ± 100 mm3) 24 or 48 h after CPA treatment and then assayed for caspase 3 activity using the fluorogenic caspase 3 substrate Ac-DEVD-AMC as described 25. Briefly, 0.5 g of tumor tissue was homogenized in 2 ml of lysis buffer (10 mM HEPES, pH 7.4, containing 2 mM EDTA, 0.1% CHAPS, 5 mM DTT, 2 mM PMSF, 10 µg/ml pepstatin, 10 µg/ml aprotinin, and 20 µg/ml leupeptin) and then centrifuged at 14,000 rpm for 20 min in a bench top Eppendorf centrifuge. The supernatant was centrifuged at 35,000 rpm for 1.5 h in a Sorvall T1270 rotor 26. The resulting supernatant (20 µg protein at 1 mg/ml) was incubated for 1 hr at 37°C with 500 µl of 50 µM Ac-DEVD-AMC in 10 mM HEPES, pH 7.4, containing 2 mM EDTA, 0.1% CHAPS and 5 mM DTT. The caspase 3 component of this activity was determined in a parallel set of assays carried out after the supernatant fraction was preincubated for 15 min at room temperature with 5 µl of the caspase 3 inhibitor Casputin. Fluorescence was measured (excitation 380 nm, emission 460 nm) using a Shimadzu model RF-1501 spectrofluorophotometer.
4-OH-CPA levels in blood, liver, and tumor were determined as described previously 22. Liver and tumor homogenates were prepared as described above. Blood samples (50 µl) were diluted to 500 µl with 0.1 M KPi buffer pH 7.4 containing 0.1 mM EDTA and 5 mM semicarbazide. Blood and tissue samples (500 µl) were deproteinized by the sequential addition of 250 µl of 5.5% zinc sulfate and 250 µl of saturated barium hydroxide. Acrolein formed during from the chemical decomposition of 4-OH-CPA was derivatized to 7-hydroxyquinoline and analyzed by HPLC. A standard curve was generated from 4-hydroperoxy-CPA (0–40 µM) dissolved in KPi buffer and processed in parallel (limit of detection (twice background), 0.3 nmol/g tissue). Integrated peak areas determined by Millennium32 software (Waters Corpn., Milford, MA) were then converted to units of nmol 4-OH-CPA produced per g tissue.
A simple noncompartmental model was used to calculate values for area under the curve (AUCINFobserved), t1/2, Cmax, and Tmax for 4-OH-CPA in blood, liver and tumor using WinNonLin software version 1.5 (Scientific Consulting, Inc., Apex, NC). 4-OH-CPA levels in tissue samples derived from CPA-treated mice were corrected for background activity (i.e., endogenous acrolein levels) based on the activity measured in the corresponding tissue samples obtained from untreated mice. To calculate pharmacokinetic parameters, the time course data for each tissue was randomly assigned into three separate data sets, each comprised of a full set of data points obtained for one of the three mice killed at each time point. The Descriptive Statistics module of WinNonLin software was used to calculate values for AUCINFobserved, t1/2, Cmax, and Tmax on the final parameters table in the form of mean values ± SEM. Statistical comparisons using a nonparametric t test were performed using Prism software version 4 (GraphPad Software, San Diego, CA).
9L/2B11 cells were grown and implanted s.c. in ICR/Fox male scid mice as described above. Tumors were grown to a size of ~500 mm3, at which time the mice were given two CPA injections at 150 mg/kg BW spaced 24 hr apart. Mice were divided into three groups (n = 5 mice and n = 10 tumors per group). One group of mice was administered with CPA i.p. (25 mg/ml CPA stock in 0.2% NaCl; systemic delivery). The other groups of mice were injected CPA, i.t., in a 23% solution of F127 prepared in 0.2% NaCl, using a syringe pump with 30-gauge needle at 1 µl/sec as described above. CPA was dissolved in the 23% F127/0.2% NaCl vehicle at concentrations ranging from 18.7 to 21.4 mg CPA/ml based on the weight of each mouse to insure an equal dose of CPA and of F127 polymer per mouse. Thus, for a 30 g mouse, a dose of 150 mg CPA/kg BW (i.e., 4.5 mg CPA/30 g mouse) was achieved by dissolving 20 mg of CPA·H2O in 1 ml of 23% F127/0.2% NaCl solution, whereas for a 32 g mouse, 21.4 mg of CPA·H2O was dissolved in 1 ml of 23% F127/0.2% NaCl solution. In both cases a total of 240 µl of the CPA/F127/NaCl solution was injected i.t. (6 × 20 µl injected into each tumor). Control mice were injected with 240 µl of 0.2% NaCl. Tumor sizes (length L and width W) were measured twice a week using an electronic digital caliper (VWR international, Marlboro, MA). A second CPA treatment cycle was administered when the tumors reached a size of ~800 mm3, i.e., day 35 after the first CPA treatment. Tumor vols were calculated using the formula: volume = π/6 (L × W) 3/2. Percent tumor regression was calculated as 100 × (V1–V2)/V1, where V1 is the tumor vol on the day of drug treatment and V2 is the vol corresponding to the maximum observed decrease in tumor size following drug treatment. Tumor doubling time was calculated as the period of time required for tumors to double in vol after the second drug treatment 27.
Scid mice bearing 9L/2B6 or P450-deficient 9L control tumors were treated with CPA and killed 24 or 48 h later. Tumor homogenates were prepared and assayed for the activity of caspase 3, an effector caspase that is activated in CPA-treated 9L tumor cells 25. CPA induced a significantly greater increase in caspase 3 activity in the P450 2B6-expressing tumors than in the P450-deficient tumors (Fig. 1). This differential response can be attributed to the activation of CPA locally, within the 9L/2B6 tumor cells, and is consistent with the much stronger anti-tumor effect seen upon CPA treatment of 9L/2B6 tumors compared to 9L tumors 17, 28.
Next we investigated whether the P450-dependent increase in CPA-induced apoptosis seen in 9L/2B6 tumors is associated with increased intratumoral exposure to 4-OH-CPA, the active metabolite of CPA. Scid mice bearing 9L and 9L/2B6 tumors were treated with CPA by i.p. injection and killed 15 min later. 4-OH-CPA level were determined in blood, liver and tumor tissue. Blood levels of 4-OH-CPA were significantly higher than those found in the liver in both 9L and 9L/2B6 tumors, indicating rapid clearance of this primary metabolite from its site of production in the liver (Fig. 2A). Surprisingly, intratumoral 4-OH-CPA levels were the same in 9L/2B6 tumors as in 9L tumors, indicating that the major fraction of tumor-associated metabolite is formed extratumorally, i.e., is derived from hepatic metabolism. A similar pattern was seen 60 min after drug administration (data not shown). 9L tumors expressing P450 2B1, which is a more active CPA 4-hydroxylase than P450 2B6 29, also showed lower 4-OH-CPA levels in tumor compared to blood and liver (Fig. 2B). A similar result was obtained in 9L/2B1 tumors treated with IFA, an isomer of CPA that is also activated by 4-hydroxylation (Fig. 2B). We conclude that the levels of P450 2B6 and P450 2B1 expressed in these tumors, while very effective at enhancing CPA’s anti-tumor activity 24, are too low to measurably increase the 4-hydroxy metabolite above the high background level that results from hepatic P450 metabolism.
The low Km of P450 2B11 for CPA (~70 µM) translates into superior anti-tumor activity in vitro and in preclinical P450 gene therapy models 24. Given this low Km, the therapeutic potential of P450 2B11 is expected to be manifest at CPA concentrations that are comparatively low (≤100 µM), as they are in CPA-treated cancer patients 30, 31. Indeed, whereas tumor-associated 4-OH-CPA levels were lower in 9L/2B1 and 9L/2B6 tumors than in liver (Fig. 2), intratumoral 4-OH-CPA concentrations were at least as high, if not higher than in liver in the case of CPA-treated 9L/2B11 tumors (Fig. 3A). Moreover, intratumoral 4-OH-CPA concentrations were significantly higher in 9L/2B11 tumors than in 9L tumors over a range of CPA doses (Fig. 3B), consistent with the occurrence of substantial intratumoral CPA activation. Intratumoral CPA activation also occurs in the case of the high Km enzymes P450 2B6 and P450 2B1, as evidenced by the enhanced apoptotic response (Fig. 1) and anti-tumor effect compared to P450-deficient tumors 24, but could not be discerned in the context of the high background level of hepatic P450 metabolism. The P450 2B11-dependent increase in intratumoral 4-OH-CPA was dose-dependent, without reaching saturation at 140 mg CPA/kg BW (Fig. 3C). The inability to saturate the active site of P450 2B11 at this dose, which corresponds to a Cmax of ~200 µM 32, suggests there is restricted uptake of circulating CPA by the tumor cells, reducing the availability of CPA for intratumoral metabolism.
More detailed pharmacokinetic analyses were carried out to determine how the low Km of P450 2B11 impacts the net exposure of tumor and host tissues to activated CPA. In 9L tumors, which do not metabolize CPA, peak concentrations of 4-OH-CPA were detected in blood and liver as early as 6 min after i.p. CPA injection, while tumor 4-OH-CPA levels reached their peak at the 15 min time point (Fig. 4A). This difference in Tmax indicates delayed diffusion of 4-OH-CPA from systemic circulation into the tumor. Moreover, at the 6 min time point, the tumor content of 4-OH-CPA was only 40% of that of blood (17.6 vs. 45.4 nmol/g tissue; Fig. 4A). In contrast, in mice bearing 9L/2B11 tumors, 4-OH-CPA concentrations were higher in tumor than in blood or liver at all times after i.p. CPA administration, except at 6 min, where similar levels of 4-OH-CPA were found in blood and tumor (Fig. 4B). Pharmacokinetic analysis revealed that, following i.p. CPA treatment, the Cmax and AUC of 4-OH-CPA were both significantly higher in 9L/2B11 tumors than in 9L tumors (Table 1). Collectively, these findings provide strong support for the occurrence of substantial intratumoral CPA activation catalyzed by the tumor cell-expressed P450 2B11.
The impact of localized, intratumoral delivery of CPA on the net exposure of 9L/2B11 tumors, and of host tissue, to 4-OH-CPA was investigated. Intratumoral CPA injection increased the Cmax and AUC values for tumor exposure to 4-OH-CPA by 1.8-fold and 2.7-fold, respectively, compared to i.p. CPA treatment (Table 1; 86.2 vs. 48.4 nmol/g tissue for Cmax; and 60.2 vs. 22.4 h.nmol/g tissue for AUC). Thus, intratumoral delivery of CPA increases the availability of CPA for metabolism by the tumor cell-expressed P450 2B11. The enhanced intratumoral metabolism of CPA was accompanied by a significant decrease in Cmax of 4-OH-CPA in both blood and liver, consistent with a shift in CPA metabolism from the liver to the tumor (Fig. 4C vs. Fig. 4B; Table 1). However, no significant change in the AUC of 4-OH-CPA was observed in either blood or liver when i.t. CPA and i.p. CPA were compared.
Next, we investigated whether further increases in intratumoral CPA 4-hydroxylation could be achieved using the slow release polymer F127 to increase the residence time of CPA in the tumor, and thereby increase the likelihood that the prodrug will be metabolized by the tumor cell-expressed P450, i.e., before it diffuses out from the tumor and into systemic circulation. F127 is soluble at 4°C but increases in viscosity and forms a gel when the temperature is raised to 37°C, as occurs following i.t. injection 33. Intratumoral delivery of CPA using a 23% solution of F127 significantly increased the AUC of tumor-associated 4-OH-CPA when compared to i.t. injection without polymer, and significantly decreased the Cmax of 4-OH-CPA in both blood and liver (Fig. 4D vs. Fig. 4C; Table 1). The lower Cmax in blood suggests there is reduced exposure of other organs to activated CPA. Both the t1/2 and the AUC of 4-OH-CPA in liver were increased by i.t. injection of CPA with polymer as compared to i.t. CPA injection without polymer or to i.p. CPA injection (Table 1). These findings suggest that the slow release of CPA from its intratumoral delivery site decreases hepatic CPA levels, which in turn leads to metabolism of CPA over an extended period of time.
We investigated the impact on anti-tumor activity of the increased exposure of 9L/2B11 tumors to 4-OH-CPA achieved by using F127 for i.t. CPA delivery. Scid mice bearing 9L/2B11 tumors were treated with CPA using a maximally tolerated dose schedule (MTD, 2 × 150 mg CPA/kg BW). CPA was administered either by i.p. injection or by i.t. injection using F127 as the vehicle. 9L/2B11 tumor growth was halted immediately in both CPA treatment groups (Fig. 5A). However, the mice treated with i.t. CPA/F127 showed significant tumor regression beginning on day 11, whereas tumor regression was not observed in the mice treated with i.p. CPA until day 21 post-CPA treatment. Moreover, the maximal tumor regression was significantly higher in the i.t. CPA/F127 treatment group (87% vs. 53% regression; p<0.0005). A second cycle of CPA treatment at day 35 effected substantial tumor growth delay but not tumor regression in either group, perhaps reflecting a preferential killing of P450 2B11-expressing tumor cells in the first treatment cycle and indicating a need for multiple rounds of P450 gene delivery in vivo for effective implementation of this strategy in the clinic. CPA-induced body weight loss was more pronounced in the mice receiving CPA via the F127 vehicle, suggesting some intrinsic toxicity of this polymer (Fig. 5B).
The efficacy of P450-based gene therapy, like that of other prodrug activation strategies, depends on three key factors: 1) the effectiveness of gene delivery in vivo; 2) the catalytic efficiency of the prodrug activation system; and 3) the biochemical and physicochemical factors that govern the metabolism, pharmacokinetics and cellular responses to the prodrug and its activated metabolites. Recent advances in vector technology have facilitated tumor-selective delivery of prodrug-activation genes and their controlled propagation as regulated by tumor-specific genetic factors and the tumor’s physiological state 34–36. Significant technical limitations in gene transfer efficiency remain, however, but can in part be compensated for by mutations that increase the catalytic activity or efficiency of the prodrug activation gene 37–39. In the case of cytochrome P450, the low catalytic efficiency of the high Km CPA 4-hydroxylase enzymes P450 2B1 and P450 2B6 toward the anti-cancer prodrug substrate CPA (Km ~ 500–1500 µM) can be greatly improved by the use of P450 2B11, a low Km CPA 4-hydroxylase (Km ~ 70 µM) 24. Presently, we used scid mice bearing 9L gliosarcomas transduced with retrovirus encoding either high Km or low Km CPA-activating P450 enzymes as a model system to characterize the metabolic and pharmacokinetic factors that underlie this improved activity and to investigate the impact of intratumoral CPA delivery using a slow release polymer.
Tissue concentrations of 4-OH-CPA, the primary activated metabolite of CPA, were lower in tumor than in blood and liver following CPA treatment of mice bearing 9L/2B6 and 9L/2B1 tumors. Moreover, intratumoral 4-OH-CPA levels in 9L/2B6 and 9L/2B1 tumors were indistinguishable from those in P450-deficient 9L tumors. A similar observation was made in the case of human glioma xenografts infected with Herpes virus encoding P450 2B1 40. Thus, in mice with tumors expressing P450 2B6 or P450 2B1, the vast majority of tumor-associated 4-OH-CPA is still derived from the liver, which is much larger in size and has a higher content of cytochrome P450 than the P450-expressing tumors. Nevertheless, 9L/2B6 and 9L/2B1 tumors both display enhanced CPA chemosensitivity 13, 28 and enhanced apoptotic responses compared to control 9L tumors, as was shown for CPA-treated 9L/2B6 tumors (caspase 3 activity; Fig. 1). This suggests that while liver-derived 4-OH-CPA may readily enter blood vessels associated with 9L tumors, it has only limited access to the relevant tumor cell compartments. As a consequence, liver-derived 4-OH-CPA is apparently much less effective at inducing tumor cell apoptosis and an overall anti-tumor response when compared to a much lower level of 4-OH-CPA formed directly within tumor cells via intratumoral P450 metabolism. Of note, 4-OH-CPA undergoes chemical decomposition to release phosphoramide mustard, a DNA crosslinking agent and the ultimate therapeutic metabolite, which unlike 4-OH-CPA, has restricted cell membrane permeability 41. Accordingly, any 4-OH-CPA molecules that decompose to phosphoramide mustard within tumor-associated blood vessels, or within the initial layers of tumor cells adjacent to the tumor vasculature, are unlikely to induce widespread tumor cell cytotoxic responses. Circulating 4-OH-CPA itself also appears to have restricted access to 9L tumors, as indicated by the 4-OH-CPA levels being consistently lower in the 9L tumors than in blood or liver, despite the fact that 9L tumors are well vascularized 42.
In contrast to 9L/2B6 and 9L/2B1 tumors, whose catalysis of 4-OH-CPA formation was undetectable in the context of the high background of liver-derived 4-OH-CPA, intratumoral formation of 4-OH-CPA by 9L/2B11 tumors was readily evident following systemic CPA administration and was associated with 4-OH-CPA levels significantly higher than in blood and liver. The high level of intratumoral CPA activation thus achieved in 9L/2B11 tumors is likely to be key to the substantially improved CPA anti-tumor response displayed in gene therapy models using P450 2B11 as compared to P450 2B6 or P450 2B1 24. Together, these findings provide proof-of-principle for the use of low Km prodrug-activation enzymes, such as P450 2B11 and improved variants 43, to augment intratumoral prodrug activation at pharmacologically relevant prodrug dosages. These findings will need to be verified using viral or other gene therapy vectors to deliver P450 genes to solid tumors in vivo, where P450 protein expression is likely to be much realized in a much smaller fraction of tumor cells than the up to 50% tumor cell coverage seen here.
Physiological conditions associated with solid tumors often lead to poor drug uptake 44, 45, which in the case of prodrug activation gene therapy may translate into inefficient prodrug activation. Presently, 4-OH-CPA production by P450 2B11-expressing 9L tumors was not saturated at a prodrug dose corresponding to a Cmax (CPA) of ~200 µM, i.e., ~ 3 times the observed Km of P450 2B11, suggesting inefficient penetration of CPA, despite the good vascularity of 9L tumors. To circumvent this problem, CPA was administered by direct intratumoral injection, which significantly improved intratumoral 4-OH-CPA pharmacokinetics, as evidenced by a 1.8-fold increase in Cmax and a 2.7-fold increase in AUC compared to systemic CPA treatment. Further improvements were obtained using the slow release polymer poloxamer 407 (F127) as vehicle for CPA delivery. This biocompatible polymer dissolves in aqueous solution at 4°C but rapidly forms a gel upon shifting the temperature to 37°C and can be used to effect slow release of lipophilic drugs and other small molecules as well as large particles such as viruses 23, 33, 46. Overall exposure of the tumor mass to 4-OH-CPA was increased 3.9-fold by i.t. CPA delivery via the slow release polymer, as indicated by AUC, and anti-tumor activity was correspondingly enhanced, as indicated by early onset and more extensive tumor regression. The effective increase in tumor cell exposure to 4-OH-CPA is likely to be even greater than the measured 3.9-fold increase in AUC, given that a substantial fraction of the AUC (4-OH-CPA) under conditions of i.p. CPA treatment is liver-derived 4-OH-CPA, which as discussed above, has poor access to the relevant tumor cell compartment. Peak host tissue 4-OH-CPA levels were also reduced with i.t. CPA delivery, both with and without the slow release polymer, and this may translate into a reduction of toxicity to sensitive host tissues. Localized chemotherapy may thus be used to shift the partitioning of P450 prodrug from the liver to the tumor. This general strategy may be implemented in a variety of ways, including using other slow release polymers that may be better optimized for small hydrophilic drugs such as CPA 40 or may be more suitable for clinical studies, or by localized instillation of prodrug to the tumor vasculature, as exemplified by preclinical and clinical studies of P450 prodrug gene therapy in pancreatic cancer 21, 47.
Grant support: NIH CA49248 (to DJW).