Introduction

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There has been a long-standing interest in the local delivery of pharmaceutical and biological agents into the peritoneum for the treatment of peritoneal diseases (Weisberger et al., 1955; Dedrick et al., 1978; Parker et al., 1981a; Markman et al., 1992; Schneider, 1994; Markman, 2001; De Bree et al., 2002). The basic goal of this treatment approach is to increase either the local drug concentration or the duration of drug exposure to a peritoneal disease process while decreasing systemic drug toxicity. Although several recent investigations have examined the efficacy of intraperitoneally delivered antibiotics, therapeutic radionuclides, and genes (Kresta et al., 1993; Meredith et al., 1996; Reimer et al., 1999), the majority of the research in this area has been conducted with intraperitoneally administered chemotherapeutic agents for the treatment of peritoneal carcinomatosis and ovarian cancer (Howell et al., 1991; Markman, 1998). Studies with chemotherapeutic agents administered intraperitoneally have yielded encouraging results for the treatment of ovarian cancer (Alberts et al., 1996; Markman, 2001). A recent consensus statement from specialists in the field of ovarian cancer recommends that intraperitoneal therapy with chemotherapy and/or biological agents be pursued as a legitimate area of research (Alberts et al., 1996; Berek et al., 1999). Reevaluation of the role of intraperitoneal chemotherapy for treatment of ovarian cancer has also recently been suggested (Kaye, 2001).

The normal pathway of drug clearance from the peritoneum is either through direct absorption across the peritoneal membrane or by drainage into the lymphatic system through absorption by the diaphragmatic stomata (Zakaria et al., 1996). Although most intraperitoneally delivered drugs are rapidly cleared from the peritoneal fluid, this method of administration can achieve much higher peak concentrations in the peritoneal fluid compared with the same drug administered intravenously (20-fold higher for cisplatin and carboplatin to as high as 1000-fold for Taxol) (Howell et al., 1991; Markman et al., 1992). Although these drug levels quickly equilibrate with plasma after termination of the peritoneal infusion, transiently elevated peritoneal drug levels provide a significant therapeutic advantage. Unfortunately, this rapid clearance from the peritoneum counteracts the advantages derived from intraperitoneal infusion. One approach to improve peritoneal drug delivery is to encapsulate the drug in a liposome (Parker et al., 1981b; Rosa and Clementi, 1983).

Liposomes are spontaneously forming lipid spheres that have been designed to encapsulate a variety of pharmacological agents (Lasic, 1996). By encapsulating a drug in a liposome, pharmacokinetics and volume of distribution of the drug can be greatly altered without diminishing its therapeutic efficacy. Intraperitoneal administration of a drug encapsulated within a liposome effectively shifts the drug clearance pathway away from direct absorption through the peritoneal membrane into clearance by lymphatic drainage. This slows the clearance of liposome encapsulated drugs from the peritoneum compared with the free drug and has been shown to prolong the contact of tumor cells with drugs, increasing antitumor activity (Sadzuka et al., 2002).

Although intraperitoneally administered conventional liposomes are more slowly cleared from the peritoneal fluid than free drug, the clearance of liposomes from the peritoneum is still fairly rapid (peritoneal clearance half-life of 1-6 h) (Parker et al., 1981b; Sadzuka et al., 2000), and the retention of liposomes in lymph nodes receiving lymph draining from the peritoneum is relatively low [percentage of the injected dose <1% (% ID) per lymph node] (Parker et al., 1981b). This low lymph node retention occurs because the majority of intraperitoneally administered liposomes returns to the systemic circulation by passing through abdominal and mediastinal lymph nodes. This minimal retention of conventional liposomes in lymph nodes has been described previously for liposomes administered subcutaneously (Oussoren et al., 1997; Phillips et al., 2001a).

We have recently described a novel method of greatly increasing the retention of subcutaneously injected liposomes in lymph nodes (Phillips et al., 2000, 2001b). This method increases the retention of liposomes in the primary lymph nodes from 1 to 2 to 12%. This method consists of liposomes coated with biotin that are subcutaneously injected, followed by an adjacent injection of avidin. The avidin causes aggregation of the liposomes, which become entrapped in the next encountered lymph node. In this article, we extend this basic methodology to peritoneal drug delivery as a means of prolonging the retention of liposome-encapsulated drugs in the peritoneum while greatly increasing their deposition in lymph nodes that receive lymphatic drainage from the peritoneum. This study was conducted with biotin-coated liposomes that encapsulated blue dye and were labeled with technetium-99m (^99mTc) to enable both visual identification of lymph nodes as well as scintigraphic imaging and quantitation of liposome distribution.

	Materials and Methods

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Liposome Preparation and Characterization. Liposomes were comprised of a 50.5:45:2.5:2 M ratio (total lipid) of distearoyl phosphatidylcholine (Avanti Polar Lipids, Birmingham, AL)/cholesterol (Calbiochem, San Diego, CA)/N-biotinoyl distearoyl phosphoethanolamine (Northern Lipids; Vancouver, BC, Canada)/-tocopherol (Aldrich Chemical Co., Milwaukee, WI). Liposomes were prepared as described previously (Phillips et al., 2001b) in a laminar flow hood using aseptic conditions. A dried film was formed by mixing the lipid ingredients in chloroform and then removing the chloroform by rotary evaporation and vacuum desiccation for at least 4 h. The dried lipid film was rehydrated in 300 mM sucrose (Sigma-Aldrich, St. Louis, MO) in sterile water at a total lipid concentration of 120 µmol/ml and lyophilized overnight. The resultant lyophilized powder was then rehydrated with 200 mM reduced glutathione (GSH) (Sigma-Aldrich) and 10 mg/ml patent blue violet dye (CI 42045; Sigma-Aldrich) in Dulbecco's phosphate-buffered saline (PBS), pH 6.3, at a final total lipid concentration of 120 µmol/ml. Immediately before extrusion, the lipid suspension was diluted to 40 µmol/ml with 100 mM GSH and 10 mg/ml blue dye in PBS, pH 6.3, containing 150 mM sucrose, and extruded through a series (2 µm, two passes; 400 nm, two passes; and 100 nm, five passes) of polycarbonate filters (Lipex, Vancouver, BC, Canada) at 55°C. Extruded liposomes were washed three times in PBS, pH 6.3, containing 75 mM sucrose and centrifuged at 45,000 rpm for 45 min in an ultracentrifuge (Ti60 rotor; Beckman Coulter, Inc., Fullerton, CA) to remove any unencapsulated sucrose, GSH, and blue dye. The final liposome pellet was reconstituted in 300 mM sucrose/PBS to a total lipid concentration of approximately 60 µmol/ml and stored at 4°C until needed.

3

Liposome Labeling. Liposomes were labeled with ^99mTc as described previously (Phillips et al., 1992). A commercial kit of the lipophilic chelator hexamethylpropyleneamine oxime (HMPAO, Ceretec; Amersham Health, Princeton, NJ) was reconstituted with 5 ml of saline containing 370 MBq of ^99mTc-pertechnetate. The kits were checked for the percentage of lipophilic HMPAO using the three-step paper chromatography system as outlined in package insert. An aliquot (1 ml) of ^99mTc-HMPAO was added to a concentrated suspension of liposomes encapsulating GSH and blue dye (1 ml; phospholipid concentration 29 mM), and incubated at room temperature for 30 min. Labeling efficiencies were determined from the ^99mTc activity associated with the ^99mTc-liposomes before and after Sephadex G-25 column separation with a dose calibrator (model Mark 5; Radix, Houston, TX). For three separate labeling experiments, the labeling efficiency was 92.8 ± 2.4%.

Imaging Studies. All animal studies were conducted under the National Institutes of Health Animal Use and Care guidelines and approved by our Institutional Animal Care Committee. Imaging studies were performed on two groups of male Sprague-Dawley rats (200-300 g), experimental (n = 5) and control (n = 5). Rats were anesthetized with ketamine/xylazine (both from Vedco, St. Joseph, MO) (50:10 mg/kg, v/v) in the thigh muscle. An aliquot (1 ml) of ^99mTc-blue-biotin-liposomes was diluted with 1 ml of saline, and this volume (2 ml; 32.7 mg of phospholipid/kg; 43 MBq) was injected into the peritoneum in both experimental and control rats. The volume of liposomes chosen for this study was determined from pilot experiments. Without dilution of the liposomes, we observed less movement into the lymphatics and more liposome aggregates in the abdomen. Anterior whole body dynamic (1 min) images were acquired with rats in a prone position lying on top of the camera face (64 × 64 word image matrix with a zoom of 1.66) using a Dyna 4 gamma camera (Picker, Cleveland, OH) interfaced to a Pinnacle computer (Medasys, Miami, FL) for 60 min after administration of the ^99mTc-liposomes. Thirty minutes after administration of the ^99mTc-blue-biotin-liposomes, the experimental rats were administered avidin (5 mg in 1 ml of saline) (Sigma-Aldrich) intraperitoneally. The timing of the avidin injection was determined from previous pilot studies. At the end of 1 h, the rats were allowed to recover from anesthesia and housed for the night. At 24 h, animals were again anesthetized as described previously, and anterior static images were acquired for 1 min in a 64 × 64 image matrix.

Image Analysis. A region of interest was placed over the whole animal on the 1-min baseline image and on the 24-h static image to determine the percentage of activity that had cleared from the body of each rat through urine or feces. The 24-h counts were decay corrected and the percentage of baseline counts remaining in the animal was calculated.

Biodistribution Studies. After the imaging study, rats were euthanized by cervical dislocation. Tissues were harvested, weighed, and counted for radioactivity (multichannel analyzer; Packard Bioscience, Meridan, CT). The % ID per organ was calculated by comparison with a standard aliquot of ^99mTc-blue-biotin-liposomes.

Statistical Analysis. Values are reported as mean ± S.E. Statistical analysis was performed using Excel (Microsoft, Redmond, WA) software for a MacIntosh computer (Apple, Cupertino, CA). Statistical differences in the % ID per organ between the experimental and control groups were determined using an unpaired Student's t test. The acceptable probability for a significant difference between means was P < 0.05.

	Results

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Figure 1A shows the 24-h whole body scintigraphic images of four different experimental rats (top images) compared with four different control rats (bottom images). Figure 1B shows an enlarged image of an experimental rat compared with a control rat. These images clearly demonstrate the very different biodistributions between the experimental and the control animals. Also, the increased accumulation of ^99mTc activity in the abdominal nodes and mediastinal nodes of experimental rats is evident. All the control animals have similar organ distributions with a high concentration of activity in the spleen, which is the organ with the greatest liposome uptake in the control animals. Experimental rats given avidin have virtually no uptake in the spleen. In experimental rats, most of the activity was confined to the abdominal region, although the distribution on the images was more variable than in the control animals. From the images, it can be seen that all experimental animals had activity in the mediastinal nodes. Image analysis of the whole animal body revealed that 81.9 ± 1.5% of the dose was still retained in the body of the experimental animals compared with only 72.8 ± 0.20% retained in the body of control animals after 24 h (P = 0.001), consistent with a faster clearance from the body of the ^99mTc in the control animals compared with the rats administered avidin.

View larger version (15K): [in this window] [in a new window]   Fig. 1.   A, scintigraphic images of four experimental rats (top row) with avidin and four control rats (bottom row) acquired 24 h postinjection. Note the very different distributions in the experimental animals, which have much greater activity in the peritoneal region and in the abdominal and mediastinal lymph nodes, without activity visualized in the spleen. The control animals have consistently high spleen uptake with virtually no activity remaining in the peritoneum. B, single enlarged scintigraphic images of an experimental rat (avidin) compared with a control rat for a more detailed comparison.

The tissue biodistribution results are shown in Table 1. Blood activity is significantly higher (P < 0.001) in the control group (14.0 ± 1.7%) compared with the experimental group (0.17 ± 0.03%). The % ID measured in the spleen is also significantly higher (P < 0.01) for the control group (23.3 ± 3.9%) compared with the experimental group (0.78 ± 0.8%). Significant amounts of activity were also detected in blue-stained abdominal nodes (4.7%) and blue-stained mediastinal nodes (2.3%) in the experimental animals (Table 1). Photographs of the mediastinum and abdomen in an experimental animal at 24-h necropsy after ^99mTc-blue-biotin-liposome administration are shown in Fig. 2, A and B. The blue staining of the mediastinal nodes and the abdominal node is clearly demonstrated in these photos. No blue-stained nodes were detectable in the control animals. The activities in the kidneys and lungs were also significantly higher in the control animals compared with the experimental animals. The only organ with substantial uptake in the experimental animals was the liver, which had 7.7% ID compared with 9.8% ID in the control animals. The total activity in the blood and major organs (liver, spleen, kidneys, lungs, and blood) was much greater in the control animals compared with the experimental animals (51.7 versus 9.6%), indicating that the ^99mTc-blue-biotin-liposomes in the absence of avidin were easily cleared from the peritoneum and associated lymph nodes.

View larger version (74K): [in this window] [in a new window]   Fig. 2.   A, photograph of mediastinal lymph nodes demonstrate the marked blue staining in experimental animals receiving avidin. Control animals had no apparent blue staining of these mediastinal nodes, even though they received the same intraperitoneal dose of 99mTc-blue-biotin liposomes. B, photograph of a blue stained abdominal lymph node in an experimental animal prior to dissection and removal. The control animal had no apparent staining of abdominal nodes.

	Discussion

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Previous studies have shown that free drugs administered into the peritoneum are rapidly cleared by absorption through the peritoneal lining. In a clinical study of free cisplatin administered by continuous hyperthermic peritoneal perfusion, only 27% of the administered cisplatin remained in the peritoneal fluid at the end of a 90-min infusion (Cho et al., 1999). Most of the cisplatin dose rapidly entered the systemic circulation by direct absorption through the peritoneal membrane.

The encapsulation of drugs in liposomes for intraperitoneal administration has several potential advantages. First, direct local toxicity of the chemotherapeutic agent may be attenuated because of encapsulation of the drug inside the protective lipid bilayer of the liposome. This is important because the dose-limiting toxicity of many intraperitoneally administered drugs is abdominal pain from direct peritoneal irritation (Markman, 2001). Liposome encapsulation has been shown to have reduced local toxicity compared with free drug when extravasated into tissue (Madhavan and Northfelt, 1995).

Second, the encapsulated drug is blocked from rapid direct absorption through the peritoneal lining, resulting in increased time for the liposome-encapsulated drug to reach tumor cells, while the encapsulated drug is slowly cleared through the lymphatics. Many studies have clearly demonstrated that the pharmacokinetics of liposome-encapsulated drugs administered intraperitoneally is very different from the same nonencapsulated drug administered intraperitoneally (Parker et al., 1981; Rosa and Clementi, 1983; Sadzuka et al., 2000). Slow removal of liposomes from the peritoneal cavity seems to provide a sustained release of drug from the liposomes into the peritoneal cavity. In one study in which liposomes encapsulating cefoxitin were administered intraperitoneally, the release of cefoxitin from the liposome complex was estimated to be well in excess of the maximum inhibitory concentration (Kresta et al., 1993). Similar findings were also described in a model of peritoneally disseminated cancer in which doxorubicin encapsulated in liposomes was considered to be slowly released in the abdominal cavity from disrupted liposomes (Sadzuka et al., 2000).

Third, increased lymph node targeting is possible because liposome-encapsulated drugs are cleared through the lymphatic vessels with at least a portion of the administered drug being deposited in the lymph nodes, where it degrades and is slowly released from the liposome in high concentration (Parker et al., 1981b; Hirano and Hunt, 1985). The avidin/biotin-liposome intraperitoneal delivery system should have the same advantages as described above for intraperitoneally delivered liposomes, with the additional enhancements of greater prolongation of intraperitoneal retention and drug release as well as increased retention and release within lymph nodes.

In the last decade, research has been performed with intraperitoneally administered liposome-encapsulated anticancer agents (Malik et al., 1991; Vadiei et al., 1992; Daoud, 1994; Sharma et al., 1996, 1997). These studies have demonstrated a significantly prolonged retention time of the liposome-encapsulated chemotherapeutic agents in the peritoneum. These studies support the hypothesis that there is a marked pharmacological advantage for the treatment of intraperitoneal malignancies by encapsulating the intraperitoneally administered chemotherapeutic agent in a liposome (Vadiei et al., 1992; Daoud, 1994). Other studies demonstrate an improved toxicity profile. For instance, encapsulation of paclitaxel in a liposome has been shown to have decreased toxicity while retaining equal efficacy for the treatment of intraperitoneal P388 leukemia (Sharma et al., 1996, 1997). It is likely that the reduced toxicity results from decreased local toxicity of encapsulated paclitaxel compared with the free drug. In humans, the dose-limiting toxicity from intraperitoneal administration of paclitaxel was severe abdominal pain, which was thought to be due to direct toxicity from either the paclitaxel or the ethanol/polyethoxylated castor oil delivery vehicle (Markman et al., 1992).

The observations from the present study suggest that the biotin-liposome/avidin methodology would enhance the reservoir-like effect observed previously for standard liposome formulations by blocking rapid lymphatic transit of liposomes from the peritoneum to the systemic circulation. The interaction of biotin-liposomes with avidin apparently results in aggregation of the liposomes in the peritoneum. This aggregation greatly alters the distribution of liposomes and seems to result in a prolonged retention of liposomes in the peritoneum as well as an increased accumulation and retention of liposomes in lymph nodes receiving drainage from the peritoneum. In our study, animals that received avidin had only a minimal percentage of the injected dose of liposomes reach the systemic circulation by 24 h as evidenced by the scintigraphic images and the low % ID found in the spleen, blood, and liver at 24 h (<9% ID). In contrast, control animals, not administered avidin, had 23% ID in the spleen, 14% ID in the blood, and 9.8% ID in the liver for a total of 47% ID in these organs at 24 h. Lymph nodes in the abdomen and in the mediastinum also had greatly increased uptake in the rats receiving avidin.

The liposome biodistribution for control animals in the present study was similar to previous reports with standard liposome formulations that were administered intraperitoneally (Ellens et al., 1981; Rosa and Clementi, 1983; Allen et al., 1993). For example, Ellens et al. (1981) reported that 19% of the intraperitoneally administered liposomes were detected in the blood at 2 h, with 7% in the liver and 4% in the spleen, indicating fairly rapid clearance of liposomes from the peritoneal cavity into the systemic circulation. In another study of intraperitoneally administered liposomes, 30% of the liposomes reached the liver by 6 h (Rosa and Clementi, 1983). Allen et al. (1993) has also demonstrated that liposomes labeled with iodine-125 and administered intraperitoneally to mice achieved peak blood levels that were very close to the situation when the same dose of liposomes were administered intravenously. After reaching the blood, the liposomes had a tissue distribution that was equal to that of intravenously injected liposomes.

It must be noted that simply making liposomes larger does not increase retention in the peritoneum or lymph nodes that receive drainage from the peritoneum. Hirono and Hunt (1985) have performed an extensive study on the effect of liposome size on their subsequent distribution after intraperitoneal administration. In their studies, 50 to 60% of the intraperitoneal dose of liposomes of varying sizes encapsulating ¹⁴C-labeled sucrose cleared from the peritoneum by 5 h in all liposomes studied. These liposomes ranged in size from 48 to 720 nm. The greatest amount of [¹⁴C]sucrose (~40%) appeared in the urine after administration of the largest liposomes. The authors speculated that the large 460- and 720-nm liposomes were unstable in the peritoneum so that they rapidly released the encapsulated [¹⁴C]sucrose. It is also unlikely that simply increasing the size of the liposomes, in and of itself, would be sufficient to result in increased peritoneal and lymph node retention because particles as large as erythrocytes readily drain from the peritoneum by passing through the lymph nodes into the bloodstream. In one study of chromium-51-labeled red blood cells injected into the peritoneal cavity of sheep, 80% of the red cells had returned to circulation by 6 h after administration (Yuan et al., 1994).

In the present study, experimental animals retained a significant portion of the liposome dose in the abdominal region and in their abdominal and mediastinal nodes. This retention of liposomes in experimental animals should result in increased release of a liposome-encapsulated drug in the peritoneal fluid and in the lymph nodes receiving lymphatic drainage from the peritoneum. Delivery of liposome-encapsulated drugs using this method should provide sustained local release of drug within the peritoneum and the lymph nodes draining the peritoneum as the liposomes degrade or become phagocytized by macrophages. This delivery system could also attenuate systemic drug toxicities by greatly reducing the rate at which a drug returns to the systemic circulation by either passage through the lymphatic vessels and lymph nodes, or through direct absorption through the peritoneal membrane.

An important potential application of liposomes that encapsulate anticancer agents is in the prophylaxis of peritoneal carcinomatosis. Because 50% of patients with malignant gastrointestinal or gynecological diseases experience peritoneal carcinomatosis shortly after local curative resection, there is a great interest in delivering intraperitoneal chemotherapy during the perioperative period. In a recent study, Hribaschek et al. (2001) found that the intraperitoneal administration of the chemotherapeutic agents cisplatin and mitomycin prevented perioperative peritoneal carcinomatosis in a rat model. The rats receiving cisplatin did, however, experience severe, local toxicity with bleeding into the peritoneum and toxic necrotic reactions of the colon. Liposomes encapsulating anticancer agents delivered by the method described in this article could potentially be used for this type of perioperative chemotherapy. The potential for treatment of micrometastasis in lymph nodes secondary to lymphatic dissemination is also great. For example, liposome retention in mediastinal lymph nodes as demonstrated in this study may be efficacious in ovarian cancer therapy as metastasis to mediastinal and other lymph nodes are not uncommon findings in ovarian cancer at autopsy (Montero et al., 2000).

The investigations in this study were in normal animals. Future studies need to be conducted in models of various disease processes. Other investigations that need to be performed include determination the best timing of the avidin injection. This timing may vary depending on the disease process being treated. For example, administration of the avidin simultaneously with the liposomes is likely to result in a significantly greater retention in the peritoneum. Other variations in timing could result in increased lymph node or diaphragm targeting of the liposomes. One significant consideration would be whether this method would have any significant advantage in an advanced disease process if the lymphatics were obstructed. With lymphatic obstruction, intraperitoneally administered liposomes would likely be retained in the peritoneum and would be unable to reach the lymph nodes so that the advantages of this delivery system would be negated. In most intraperitoneal disease processes, complete obstruction of the lymphatics is uncommon and generally occurs only in advanced disease. For example, ovarian cancer is the most common cause of malignant ascites, which is often due to lymphatic obstruction. However, in a recent study only 12.5% of women diagnosed with early stage ovarian cancer presented with ascites (Eltabbakh et al., 1999).

In summary, the intraperitoneal biotin-liposome/avidin delivery method described in this article has potential as a delivery system for the local treatment of intraperitoneal and intralymphatic disease processes by increasing the retention of drugs in the peritoneum and in the lymph nodes that receive lymphatic drainage from the peritoneum.