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Preclinical Experimental Therapeutics |
Department of Neurosurgery, Northwestern University Medical School, Chicago, IL 60210, USA (M.J.A., R.M.L.); Department of Neurology, Northwestern University Medical School, Evanston Northwestern Healthcare, Evanston, IL 60201, and Northwestern University Institute for Neuroscience, Northwestern University, Evanston, IL 60208, USA (Y.N., M.W.V., E.W.-Y.K., A.C.I., R.M.L., D.R.G.); Hungarian-Japanese EM Center, Department of Pathology, University Medical School of Debrecen, H-4014 Debrecen, Hungary (P.M.)
3 Address correspondence to Dennis R. Groothuis, Department of Neurology, Evanston Hospital, 2650 Ridge Avenue, Evanston, IL 60201 (drgroothuis{at}northwestern.edu).
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Abstract |
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Key Words: brain tumors • convection-enhanced delivery • drug delivery • glioma • interstitial fluid pressure
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Introduction |
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The prevailing consensus has been that the BBB is a limiting factor in drug delivery after systemic administration. Consequently, several different methods have been developed in an attempt to circumvent this limitation, including hyperosmotic BBB disruption (Kroll and Neuwelt, 1998; Williams et al., 1995), chemical barrier modification (Black et al., 1997), attempts to link therapeutic agents to compounds that have transporters across the BBB (Bickel et al., 2001; Zhang and Pardridge, 2001), and most recently, direct administration of drugs into and around brain tumors (Hassenbusch et al., 2002; Hau et al., 2002; Prados et al., 2002; Reardon et al., 2002; Weber and Hingorani, 2002). The last method may include the placement of drug-loaded wafers around a tumor resection bed, infusion of agents into or around a tumor resection cavity, or direct infusion of drugs into the tumor mass. The concept of direct drug administration into brain tumors is not new; in 1991, Tomita reviewed the literature up to that point and reported clinical studies of interstitial chemotherapy as far back as the early 1960s (Tomita, 1991). Since the publication by Morrison et al. (1994) about microinfusions into brain, there has been a resurgence of interest in this method for the treatment of brain tumors; it is often called convection-enhanced delivery (CED).
Regardless of the route of administration, however, the role of delivery in therapeutic effectiveness remains a confounding variable. Given contemporary methodology, and apart from whether a study is conducted in an animal model or in humans, in most studies the basis for treatment failure cannot be clearly assigned either to inadequate delivery or to lack of therapeutic effectiveness on the part of the agent being used. The experiments in this study were designed to address this issue. We hypothesized that if we had an agent that was unequivocally capable of killing tumor cells, we could identify the delivery parameters that would deliver that agent with a concentration and time sufficient to accomplish the goal: 100% tumor kill. We were unable to identify a conventional brain tumor therapeutic agent (drug, antibody, or toxin) for which we could unquestionably assume 100% effectiveness in killing brain tumor cells. To remove the issue of an agent's therapeutic effectiveness from the mix of confounding variables, we therefore decided to use formalin, which is a chemical that would kill (i.e., "fix") 100% of tissue exposed to it for a sufficient period. The selection of this agent may be controversial, but it enabled this study to begin with an agent that would unequivocally kill tumor cells, providing that delivery was adequate.
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Materials and Methods |
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In Vitro Sensitivity Testing
In vitro sensitivity of RG-2 or D54-MG cells to formalin was tested. Formalin solutions were prepared as neutral buffered formalin solution (Luna, 1968), except that MEM-A was used instead of distilled water, and the amount of formalin was varied to make a range of concentrations. A single-cell suspension of RG-2 or D54-MG cells (105 ml-1) was placed in formalin-MEM-A, in concentrations ranging from 0.15% to 10%, and in a control of MEM-A alone. Cells were placed in an oscillating shaker for exposure periods of 10 min, 20 min, or 2 h, after which they were centrifuged, washed, transferred to fresh MEM-A, and evaluated daily. Initially, we tried to use the trypan blue exclusion assay to assess viability, but the formalin affected membrane permeability and gave false-positive results, suggesting tumor cell death when the tumor cells actually were viable. Since we were unable to sort live from dead tumor cells, we resorted to a more direct measure of viability: We measured the time it took for cell populations to reach confluence.
Animal Preparation
Fisher-344 rats (Harlan Industries, Indianapolis, Ind.) were anesthetized with isoflurane/nitrous oxide/oxygen anesthesia (1.5/30/70, v/v/v) and injected with 10 µl RG-2 glioma cells (106 cells/ml) s.c. in the flank (Groothuis et al., 1983). Rats were used for experiments when the tumors measured between 5 and 8 mm in diameter. Athymic mice (nu/nu, Harlan Industries) were anesthetized and injected with 50 µl of D54-MG cells (106 cells/ml) s.c. in the flank; mice with tumors were used for experiments when the tumors measured between 5 and 8 mm in diameter.
For chronic infusions, rats were anesthetized with ketamine-xylazine. Osmotic infusion pumps (2ML1, Durect Corp., Cupertino, Calif.), loaded with either normal saline or 10% neutral buffered formalin, were inserted into a subcutaneous pocket on the back. The pumps were attached to a polyethylene catheter, which was in turn attached to a half-inch 25-gauge needle. The needle was inserted into the tumor, and two surgical ties were placed around the needle and overlying skin to hold it in place. The incision was closed, the rat was allowed to awaken, and measurements of tumor size in two perpendicular horizontal axes were recorded daily for 14 days. Sixteen tumors received an infusion of saline and 16 received an infusion of 10% formalin. Tumor volume was calculated from the formula Vt = (lw2)/2, where l and w are length and width, respectively (Bullard et al., 1981). For each tumor, the volume was normalized by the tumor volume on day 0, before treatment. At the end of the infusion, the tumor was removed and frozen; later, the tumor was serially sectioned at 20-µm thickness on a cryostat.
For acute infusions, rats or mice were anesthetized with ketamine-xylazine. Polyethylene tubing was attached to a Sage 365 programmable infusion pump (ATI Orion, Boston, Mass.). The other end of the polyethylene tubing was attached to a half-inch 25-gauge needle and inserted into the tumor as described above.
In RG-2 tumor-bearing rats, infusions of 10% neutral buffered formalin were made at the following rates and durations: 2 µl/min x 2 h (n = 12), 4 µl/min x 2 h (n = 14), 6 µl/min x 2 h (n = 5), 8 µl/min x 2 h (n = 6), 12 µl/min x 1 h (n = 5), and 48 µl/min x 15 min (n = 9). The infusion site was visualized directly; in those infusions in which fluid could be seen leaking from the skin, the needle was repositioned. After the experiment, the needle was removed, the rat was allowed to awaken, and measurements of tumor size were recorded daily for 14 days. Normalized tumor volume was calculated as described above. At the end of the infusion, the tumor was removed, frozen, and serially sectioned at 20-µm thickness on a cryostat. An additional group of tumors (n = 12) was infused with 10% formalin for 15 min at 48 µl/min, and the rats were allowed to survive for 50 days.
In the D54-MG tumor-bearing mice, infusions of 10% neutral buffered formalin were made at the following rates: 4 (n = 5), 8 (n = 5), and 24 (n = 16) µl/min. The duration of the infusion was 60 min.
Distribution of 14C-Formaldehyde
To examine the distribution of formalin within the tumors, 2 µCi 14C-formaldehyde (53 mCi/mmol, >99% radiochemical purity [Moravek Biochemicals, Brea, Calif.]), prepared in 10% neutral buffered formalin, was infused into RG-2 tumors at 2 µl/min for 2 h, and at 48 µl/min for 15 min. These rate combinations were selected because they represented either a failure of delivery (2 µl/min for 2 h [Fig. 1]) or successful delivery (48 µl/min for 15 min [Figs. 2 and 3]) in the prior experiments. At the end of the experiment, the tumors, including a margin of the tissue into which the tumor was growing, were removed and frozen (-80°C) in liquid Freon (E.I. Du Pont de Nemours, Wilmington, Del.) within 1 min of the conclusion of the experiment. Preparation of autoradiographs of the s.c. tumors has been described previously (Blasberg et al., 1981; Groothuis et al., 1983; Vriesendorp et al., 1987). The autoradiographs were digitized at a resolution of 50 µm/pixel with a video-based digitizing system, along with calibrated 14C-methyl methacrylate standards. After the autoradiographs were developed, the tissue sections were stained with hematoxylin-eosin, digitized, and aligned with the autoradiographic images in computer memory. Regional tissue measurements of radioactivity (nCi/g) were obtained from the autoradiographs by using the histological images to define the region of interest. The data were analyzed to obtain the fraction of injected 14C activity contained in different tumor regions at the termination of the experiment.
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Subcutaneous RG-2 Tumor Growth in Rats in Response to Formalin Infusion
The size of tumors at the initiation of the formalin infusions is listed in Table 1. The time course of tumor volume in the group that received saline infusions with the osmotic minipump is shown in Fig. 4A; by two weeks, this tumor volume had increased 100-fold. In contrast, tumor volume in the group that received 10% formalin by osmotic minipump for seven days had increased only sevenfold (Fig. 4B). However, each tumor in this group continued to grow during the treatment period. Histological examination of these tumors showed little difference from those receiving saline. Aside from size, the microscopic findings were similar, and tumor cells that had been killed during the formalin infusion could not be distinguished from cells that were alive at the termination of the experiment.
The groups that received acute infusions of 10% formalin at infusion rates of 2 to 48 µl/min all showed a prominent reaction to the infusion (Figs. 1 and 4), as illustrated by the time course of "apparent" tumor volume immediately following the infusions. During the first 24 h after the infusion, there was intense swelling, resulting in an increase in the apparent tumor size of up to 50 times. This acute edematous reaction subsided between 4 and 6 days after the infusion. In the group that received 2 µl/min for 2 h, the tumors displayed exponential growth by day 14 (Fig. 4C), and when examined individually, every tumor in this group was growing. In the 4-, 6-, and 8-µl/min groups, which were all infused for 2 h, some individual tumors did not have any growth by day 14, whereas others were clearly growing larger. In the 4-µl/min group, three (20%) of 14 tumors had no growth over the 14 days. In the 6-µl/min group, two (40%) of five tumors had no growth, and in the 8-µl/min group, one (20%) of five tumors had no growth. Microscopically, there was little apparent difference between these tumors and those infused with the osmotic pumps: Viable tumor cells and those effectively fixed and killed by the formalin could not reliably be distinguished.
In the 12-µl/min x 1 h group (n = 5) and in the 48-µl/min x 15 min group (n = 9), all tumor growth was suppressed. Histologically, however, the tumors appeared as described above: Formalin-fixed tumor cells could not be reliably distinguished from those that were alive at the termination of the experiment.
Because we could not microscopically distinguish live from dead tumor cells and were thus uncertain about whether the tumors had been eradicated, an additional 20 tumors were infused with 10% formalin at 48 µl/min for 15 min, and their condition was followed for 50 days. Since we wanted to have both successful and unsuccessful outcomes ("cures" and treatment failures, respectively), a spectrum of tumor sizes was used, and the mean tumor size at the time of treatment for the additional 20 animals was larger than in the original groups of animals (Table 1). Selected animals were killed at 2, 14, and 31 days after infusion and examined histologically. The remaining 10 animals were killed at 50 days (Fig. 2). Of these, five were "cures" and five were treatment failures. Following the formalin infusions, the initial course of all animals was similar; that is, by the day after the infusion, there was a large edematous area in which the tumor could not be distinguished from the reaction to the formalin. Subsequently, the edematous area decreased in volume and became firm and rubbery, but at this time (4-7 days after the infusion), the animals that would eventually be cured could not be distinguished from those that would not. By day 14, skin overlying the tumor had become necrotic and ulcerated; in animals that were tumor free, the ulcers healed completely, skin covered the site, and hair regrew. Upon microscopic examination of the tumor site in these five animals, there was a small abscess in one, granulation tissue in one, and fibrotic scars in the remaining three animals. In the group of treatment failures, the animals that had persistent tumor had a hard, firm rim around a central craterlike ulcer. The center of the ulcer was tumor free, but the rim, which was of variable thickness, had both macroscopic and microscopic tumor foci. In the animals with tumor cells, the tumors were not histologically different from untreated tumors.
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Distribution of 14C-Formaldehyde
In the RG-2 tumors that received a 2-µl/min infusion of 14C-formalin for 2 h, the distribution pattern was typically patchy, with higher concentrations of 14C activity in areas of necrosis or in tissue surrounding the tumor (Fig. 5). In each of these tumors, there were islands of viable tumor cells that did not contain any 14C activity. In contrast, in the RG-2 tumors that received an infusion of 48 µl/min of 14C-formalin for 15 min (Fig. 6), the highest concentrations of 14C activity were found at the tumor edge, but some radioactivity was present in all parts of the tumor. In some of the larger tumors, which contained islands of necrosis, 14C activity was just beginning to accumulate in the necrotic areas (Fig. 6). In both groups of tumors, greater than 95% of the administered 14C activity was found in either necrotic areas of the tumor or in the subcutaneous tissue surrounding the tumor.
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Discussion |
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The selection of formalin was only somewhat arbitrary. We could not identify an existing therapeutic agent that could confidently be expected to kill all tumor cells. Because we wanted to unequivocally separate the issue of sensitivity from that of therapy, we used 10% formalin, which we were confident would kill all cells if delivered adequately. Formalin actually consists of several different chemical species, including formaldehyde, and reacts extensively with proteins, nucleic acids, carbohydrates, and to a lesser extent with lipids (Hayat, 1981). The reactions are rapid and result in cross-links between many different compounds. Formalin may be considered a drug of high reactivity and low specificity. As to its effects as a drug, we were interested in knowing about the in vitro chemosensitivity of the tumor cells to formalin despite the poor correlation between in vitro chemosensitivity and in vivo responsiveness in general (Yung, 1989). We conducted in vitro chemosensitivity testing on the tumor cells, in part to determine the relationship between the concentration-time product of exposure and tumor cell kill. This showed that both time and concentration were important. Any concentration over 1.5% killed all tumor cells, and all concentrations lower than 1.5% delayed growth, although the cells eventually recovered and exhibited exponential growth. When 1.5% formalin was used for 20 min, growth was delayed, whereas a 60-min exposure killed all cells. Unfortunately, direct comparisons between formalin and conventional chemotherapeutic agents cannot be made. Most therapeutic agents have specific cellular targets and do not affect all cellular elements, as does formalin. The concentration-time products obtained with 10% formalin will almost certainly underestimate those required with conventional therapeutic agents, which in turn means that the infusion parameters (infusion rate and duration) used in these experiments will likely underestimate those needed for studies with conventional drugs and experimental tumors.
In light of these results, it may be important to revisit the issue of in vitro chemosensitivity from a delivery aspect. Since the present studies have shown that inadequate drug delivery can be a major reason for treatment failure, the inability to correlate in vitro and in vivo chemosensitivity and in vivo response in previous studies of this type may have been due to failed delivery. Many studies of in vitro chemosensitivity use fixed time points to assay the tumor cell responsiveness. A common time point is 48 h (Miller et al., 2002). It is very unlikely, however, that either direct infusion of drug into tumors or intravenous infusions will be able to maintain a required concentration throughout the entire tumor volume for 48 h. In vitro chemosensitivity testing may need to look at a spectrum of concentration-time products, including exposure times as short as 10 or 15 min. Some of the differential responsiveness may be the result of delivery and not sensitivity (Friedman et al., 1995).
Another potential point of controversy in these studies of RG-2 and D54-MG subcutaneous gliomas is the relevance of using a subcutaneous tumor as a model for an intracerebral tumor. The use of subcutaneous xenografts to study chemoresponsiveness is common (Friedman et al., 1995; Goudar et al., 2005; Hjortland et al., 2004). Although we have established that many of the properties of the RG-2 rat glioma are the same whether the tumor is intracerebral or subcutaneous (Molnar et al., 1999), we have not studied all of the physiological properties that would influence drug delivery. We have also used a model in which RG-2 or D54-MG cells are injected into the brain through an indwelling cannula and in some instances produced ideal tumors for CED therapy (Vavra et al., 2004). Common problems with this model, however, are that there is no convenient way to monitor tumor size and that some tumors deviate from the ideal, becoming multilobulated, and can be found growing at some distance from the end of the cannula. In addition, as an exercise in isolating delivery issues, the infusion rates used in the present study could not have been tolerated by an intracranial model. In our previous study of 14C-sucrose infusion into normal rat brain (Vavra et al., 2004), we found the maximum tolerated infusion rate to be between 0.5 µl/min for 8 h and 1 µl/min for 4 h. If an intracerebral tumor, which in and of itself causes increased intracranial pressure, were treated with CED infusion at the rates we found necessary, we would never have been able to kill any of the RG-2 tumors before the animals succumbed to increased intracranial pressure. Instead of being a drawback, therefore, the use of a subcutaneous model enabled better monitoring of tumor size, confidence that the tip of the infusion cannula was in the tumor center, and the ability to explore infusion rates that are prohibitive in intracerebral models.
If failure to kill subcutaneous gliomas in this study can be explained by the inability to maintain a minimum concentration-time product of formalin in the RG-2 gliomas, it is important to understand this issue more completely. We believe that failure to maintain the minimum concentration-time product can largely be explained by the pressure in the tumor extracellular space. Jain et al., who have measured IFP in several different tumor lines, have found IFP values as high as 50 mmHg (Gutmann et al., 1992; Kristjansen et al., 1993; Leunig et al., 1992; Netti et al., 1995; Znati et al., 1996). We have measured IFP in RG-2 gliomas and found it to be 9.1 ± 2.1 mmHg (Navalitloha et al., submitted). We have also measured the efflux rate of 14C-sucrose from RG-2 and D54-MG gliomas. In the RG-2 gliomas, the efflux half-time was 7.3 ± 0.7 min (Vavra et al., 2004). In contrast, the efflux half-time from D54-MG gliomas was 18 ± 4.2 min (unpublished observations). The distribution of 14C-formalin in the present study also indicates the rapid rate of efflux from the tumor (Fig. 6). At the end of the 48-µl/min x 15 min infusion, more than 95% of the 14C activity was in tissue surrounding the tumor, and the lowest concentration of 14C-formalin was always in the viable tumor center. There are, therefore, two consequences of local pressure gradients within tumors: (1) Areas in which the tumor IFP exceeds the infusion pressure will block drug delivery because of the pressure differential, and (2) drug will leave tumor tissue quickly because of the high efflux rates caused by the pressure gradient from within the tumor into the surrounding tissue. It is also important to emphasize that increased IFP will affect drug delivery regardless of whether the drug is administered intravenously, intra-arterially, or by direct infusion. Up until the present, therapeutic studies in animal models have usually been conducted independently of knowledge about underlying tumor physiology (Friedman et al., 1995; Giussani et al., 2002, 2003; Viola et al., 1995). Until the variability of issues surrounding delivery is understood, there may continue to be a "hit or miss" outcome to many of these experiments.
It is interesting to briefly compare the methods used in this study to those used in clinical protocols in which conventional chemotherapy drugs (Mardor et al., 2001), monoclonal antibodies (Reardon et al., 2002), viruses, or toxins linked to receptor ligands (Weber and Hingorani, 2002) are being used. Only a few of these studies have been published (Cohen et al., 2003; Laske and Rossi, 2002; Parney et al., 2005; Sampson et al., 2005). However, there are some major differences between these studies and our own. Many of the trials are infusing into a resection cavity or into brain around a resection cavity, which involves tissue pharmacokinetics that are different than those involved with infusion into a tumor. Most of these clinical trials are using infusion rates of 2 to 6 µl/min, which are lower than the lowest successful infusion rates we used in the RG-2 tumors. Finally, the human tumors being treated are extremely large in comparison to the experimental RG-2 or D54-MG tumors in the present study.
The major conclusion of this study is that, even with a potent toxin, the infusion rates needed to overcome local tissue pressure gradients are high and perhaps beyond the rates that can be used in human brain tumors. This suggests that clinical trials using direct infusion into tumors may have negative outcomes not because the agents being used are ineffective, but because the agents are not being delivered to the intended target tissue with a concentration-time product that is adequate to kill the tumor cells. These studies also suggest that the terminology that is being used in much of the literature about direct infusion, or CED, is inadequate to properly describe the pharmacokinetics of drug distribution. The term volume of distribution is often used to describe the volume of tissue reached by the infusion. However, this term ignores heterogeneity in the values of local drug concentration. Since hydrostatic pressure is the force that drives convection, the distribution of solvent and solute may reach large tissue volumes but, in the process, simply bypass local tissue regions in which the tumor IFP exceeds the pressure generated by the infusion. This may result in inhomogeneous distribution (illustrated by Fig. 5) in which the most viable tumor regions are never adequately exposed to a drug, even though they are located within the total volume in which drug is being distributed.
Several questions remain unanswered. How variable are tumor IFP and efflux time within individual tumors and for different tumors? Can the tumor IFP be modified by local or systemic maneuvers to effectively decrease the rate of efflux from tumors (Kristjansen et al., 1993; Lee et al., 1992, 1994), thereby increasing residence time of therapeutic agents as suggested by Jain et al. (Gutmann et al., 1992; Kristjansen et al., 1993; Leunig et al., 1992; Netti et al., 1995; Znati et al., 1996)? Ongoing trials of direct infusion into human tumors do not monitor the distribution process, either in terms of the volume of distribution or in terms of drug concentration. In light of the present studies, it is apparent that treatment failures are to be expected. In our studies, there were also treatment successes in individual tumors at rates lower than those required to kill all tumors, suggesting that the placement of infusion catheters may have played a role in the distribution of the administered drug or that reflux along the catheter might have occurred in some tumors. We believe that clinicians need to monitor the delivery process for each individual case to ensure that the therapeutic agent is reaching the target tumor cells in sufficient concentration and for sufficient time as to be therapeutic, unless these variables can be controlled to such an extent a priori that CED can be used as an effective therapeutic tool without such individual monitoring. We believe that the development and application of drug delivery monitoring methodologies in individual patients must be developed to improve therapeutic outcome in a tumor known for its physiologic variability.
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Footnotes |
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2 Mir J. Ali is currently with the University of California, San Francisco.
4 Abbreviations used are as follows: BBB, blood-brain barrier; CED, convection-enhanced delivery; IFP, interstitial fluid pressure; MEM, minimum essential media.
Received for publication August 8, 2005. Accepted for publication November 18, 2005.
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References |
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