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Biophysical Journal logoLink to Biophysical Journal
. 2016 May 10;110(9):2106–2119. doi: 10.1016/j.bpj.2016.03.040

Interstitial Pressure in Pancreatic Ductal Adenocarcinoma Is Dominated by a Gel-Fluid Phase

Christopher C DuFort 1, Kathleen E DelGiorno 1, Markus A Carlson 1, Ryan J Osgood 3, Chunmei Zhao 3, Zhongdong Huang 3, Curtis B Thompson 3, Robert J Connor 3, Christopher D Thanos 3, J Scott Brockenbrough 1, Paolo P Provenzano 1, Gregory I Frost 3, H Michael Shepard 3, Sunil R Hingorani 1,2,4,
PMCID: PMC4939548  PMID: 27166818

Abstract

Elevated interstitial fluid pressure can present a substantial barrier to drug delivery in solid tumors. This is particularly true of pancreatic ductal adenocarcinoma, a highly lethal disease characterized by a robust fibroinflammatory response, widespread vascular collapse, and hypoperfusion that together serve as primary mechanisms of treatment resistance. Free-fluid pressures, however, are relatively low in pancreatic ductal adenocarcinoma and cannot account for the vascular collapse. Indeed, we have shown that the overexpression and deposition in the interstitium of high-molecular-weight hyaluronan (HA) is principally responsible for generating pressures that can reach 100 mmHg through the creation of a large gel-fluid phase. By interrogating a variety of tissues, tumor types, and experimental model systems, we show that an HA-dependent fluid phase contributes substantially to pressures in many solid tumors and has been largely unappreciated heretofore. We investigated the relative contributions of both freely mobile fluid and gel fluid to interstitial fluid pressure by performing simultaneous, real-time fluid-pressure measurements with both the classical wick-in-needle method (to estimate free-fluid pressure) and a piezoelectric pressure catheter transducer (which is capable of capturing pressures associated with either phase). We demonstrate further that systemic treatment with pegylated recombinant hyaluronidase (PEGPH20) depletes interstitial HA and eliminates the gel-fluid phase. This significantly reduces interstitial pressures and leaves primarily free fluid behind, relieving the barrier to drug delivery. These findings argue that quantifying the contributions of free- and gel-fluid phases to hydraulically transmitted pressures in a given cancer will be essential to designing the most appropriate and effective strategies to overcome this important and frequently underestimated resistance mechanism.

Introduction

The delivery of fluid and nutrients to tissues is governed by well-established principles of solute transport and fluid flux (1, 2). Solute flux is driven by diffusion, which depends on concentration gradients, and convection, which is determined by a hydrostatic pressure gradient that promotes fluid efflux from vessels in opposition to an oncotic gradient favoring fluid retention (1). The very low interstitial fluid pressures (IFPs) that facilitate fluid efflux in normal organs can increase substantially in neoplasia (3). Aberrantly elevated IFPs can impede systemically administered drugs and compromise the treatment of solid tumors (4, 5, 6).

The study of IFPs in pathologic conditions began with the work of Starling and his inquiries into dropsy (edema) (1). This condition of excess free fluid in the interstitium remained a major focus for several decades and gave rise to a series of “fluid equilibration” techniques, including the needle, modified wick-in-needle (WN), perforated capsule, and micropipette (7, 8, 9, 10), that were capable of measuring free-fluid pressures. Increased fluid pressures in solid tumors were first described by Young et al. in transplanted Brown-Pearce carcinomas as a potential mechanism driving the dissemination of cancer cells (3). Fluid pressures have since been studied in a variety of in vitro, ex vivo, and engrafted tumor models and have typically ranged from 10 to 40 mmHg, comparable to the hydrostatic pressures in terminal capillaries and presenting, therefore, a relative barrier to drug delivery (11, 12, 13). Similar measurements have rarely been performed in autochthonous cancers, however, and such modestly elevated pressures could not in any case explain the profound hypoperfusion (14) and vascular collapse (6) found in pancreatic ductal adenocarcinomas (PDAs).

PDAs are characterized by a robust desmoplastic or fibroinflammatory stroma; a dense extracellular matrix, including exceedingly high concentrations of interstitial hyaluronan (HA); and a sparse and largely collapsed vasculature. In addition, the vessels that are present appear structurally intact and lack the fenestrae and increased interendothelial junctions observed in engrafted tumor models (15). Moreover, IFPs in autochthonous PDAs measured with the classic fluid equilibration techniques are paradoxically even lower than those encountered in the more artificial systems noted above.

It has long been appreciated, though more recently often overlooked, that interstitial fluid is normally found in two phases: freely mobile (unbound) and less mobile (gel-bound) fluid (16). The majority of tissue interstitial fluid resides in a viscoelastic gel-fluid phase comprised principally of HA. HA is a naturally occurring, soluble, unbranched glycosaminoglycan (GAG) composed of repeating disaccharide units of N-acetyl-D-glucosamine and D-glucuronic acid that can reach megadalton molecular masses (17). HA is found in small amounts throughout most organs of the body and at very high concentrations in the joint space and umbilical cord (18, 19). Its high negative charge contributes to its ability to imbibe and complex large amounts of water. HA can bind up to 15 molecules of water per disaccharide unit and organize the complexed water into one or more distinct bound states (20, 21). Together with lesser amounts of other GAGs and proteoglycans, HA establishes a sizable, less mobile, gel-fluid phase in tissue interstitia. These properties explain its important roles in normal physiology, such as maintaining skin turgor and underpinning the shock-absorbing capabilities of the joints (22, 23). That the preponderance of interstitial fluid is trapped in a gel-fluid, viscoelastic phase also explains why cutting into tissues does not release appreciable free fluid (8, 24, 25).

We hypothesized that the majority of interstitial fluid in PDA is also in an HA-dependent, less mobile (gel-fluid) phase and that fluid pressures in PDA have heretofore been markedly underestimated because the original methods were ill-suited to measuring pressure in this phase. In the following studies, we tested these hypotheses directly by investigating the sources of fluid pressures across a range of physiologic states and assessing the abilities of different types of instrumentation to capture these pressures. We measured IFPs in normal tissues, engrafted tumors, and autochthonous cancers with both WN, to measure free-fluid pressures, and a piezoelectric pressure catheter (PC), which we propose can capture gel-fluid pressures. These complementary methods enable dissection of the contributions of each fluid phase to the perfusion barriers generated in distinct contexts and tumor types.

Materials and Methods

Mouse strains

All animal studies were approved by Institutional Animal Care and Use Committees at Fred Hutchinson Cancer Research Center or Halozyme Therapeutics, Inc., and were conducted in accordance with the recommendations outlined in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. KrasLSL-G12D/+;Trp53LSL-R172H/+;Pdx1-Cre (KPC) mice have been described previously in detail (26, 27). Analyses were also conducted in normal organs of 2- to 4-month-old wild-type mice on a mixed 129Sv/Bl6 background and in allografted and xenografted tumors in nude mice (NCI Charles River, Frederick, MD).

Engrafted tumors

Subcutaneous tumor grafts were made by injecting nude mice with 1.5 × 106 cells per site with the following cell lines: primary murine PDA cell lines mPDA#1 and mPDA#2 (27), NIH3T3 (murine fibroblasts), HT-1080 (human fibrosarcoma), or B16-F10 (murine melanoma) (American Type Culture Collection, Manassas, VA). Peritibial tumor grafts were generated by injecting 5 × 106 control BxPC3 (human pancreas cancer) cells, or BxPC3 cells transduced to overexpress HA synthase 3 (BxPC3/HAS3), adjacent to the tibial periosteum.

Measuring IFP

PC transducer

IFP measurements were performed using a Millar Mikro-Tip PC transducer SPR-1000 probe (330 μm diameter) or SPR-671 probe (460 μm diameter) with a dynamic pressure range from −50 to 300 mmHg; the probes possess a shielded, recessed, side-mounted piezoelectric pressure sensor (Millar Instruments, Houston, TX). The catheter was connected to a PCU-2000 pressure control unit (Millar Instruments) and an ADInstruments PowerLab data acquisition system (ADInstruments, Colorado Springs, CO), as previously described (6, 28). The probe was allowed to equilibrate in water for 30 min and was then set to zero in a droplet of water. The system was calibrated to 0, 25, and 100 mmHg before each measurement per the manufacturer’s recommendations, and then to 30, 60, 90, 120, and 150 mmHg with a sphygmomanometer; calibrations performed using columns of water of varying heights yielded essentially identical standard curves. Instrument calibration was reconfirmed after each experiment (29). The PC probe was inserted into a 20- to 21-gauge needle used to guide insertion into the tissue of interest, after which the needle was withdrawn, leaving the probe in place; this procedure minimizes trauma to both tissue and probe. We ensured that the probe could be freely rotated and inserted and withdrawn within the preformed needle track without resistance or shearing of tissue to confirm the lack of adherence of or interference from solid-tissue elements to the side-port sensor. Real-time pressure data were recorded and analyzed using LabChart software (ADInstruments).

WN technique

WN measurements were performed with minor modifications to the original method of Scholander et al. (8) and further refined by Fadnes (9). WNs were generated as previously described (9); several needles were also provided by the Steele Laboratory (Harvard University, Cambridge, MA). For WN measurements, needles containing three to five nylon sutures were filled with a dilute heparin solution, equilibrated for 30 min in water, and connected to a disposable transducer linked to a bridge amplifier (ADInstruments) and a PowerLab data acquisition system. Calibration was performed before every experiment by either of the two methods described above. After tissue insertion, the connecting tubing of the WN was pinched and released to confirm the appropriate upward and downward pressure deflections reflecting uninterrupted communication with free fluid in the tissue (9).

Intravital IFP measurements

Mice were anesthetized with continuous-inhalation isofluorane (2.5%, 1 L/min). Subcutaneous IFP measurements were made under the skin over the abdomen. Muscle IFP measurements were performed in the right gastrocnemius muscle. For liver, pancreas, and autochthonous PDA measurements, mice underwent laparotomy and the probe was inserted under direct visualization.

In vitro experiments

Sealed Vacutainers (Becton Dickinson, Franklin Lakes, NJ) were used to simulate a closed in vitro system. After calibration, the WN was inserted through the rubber stopper. A 21-gauge needle was then inserted through the stopper, the PC was threaded through, and the needle was withdrawn, leaving the two probes adjacent and at equal heights. Aliquots of pure water or a concentrated HA solution were then added in distinct sequences. For example, 1 mL of water or 1 mL of HA in solution (concentration between 0.5% and 3%) was first added to the Vacutainer and the tube was resealed. A 25-gauge needle was inserted through the stopper, allowing any air pressure to be released, and the tube was inverted. Probes were allowed to equilibrate in the liquid for ∼30 min to ensure a zeroed reading before the subsequent injections. For experiments involving mixing, a 3 mL syringe containing HA or water was inserted through the stopper, and above the line of initial liquid, and a second, empty 3 mL syringe was inserted through the stopper and remained in the bottom of the fluid. After each injection, the syringe plunger was withdrawn and then quickly depressed several times to mix together the solutions, while keeping the fluid-filled syringe from refilling due to increased pressure in the Vacutainer.

Cardiac cessation

Mice were anesthetized as above, and the PC and WN were placed in the appropriate tissue. Isoflurane (5%, 1 L/min) was administered for 5 min to induce respiratory arrest. Cardiac arrest was confirmed visually. A subset of mice were instead cervically dislocated and then rapidly decapitated to induce arrest by a different means.

Histology and histochemistry

Harvested tissue and tumor specimens were fixed in 10% neutral buffered formalin, embedded in paraffin, and sectioned (5 μm). Sections were stained with hematoxylin and eosin (H&E) (VWR, Radnor, PA), Masson’s Trichrome (IHC World, Woodstock, MD) to highlight total collagen content, and Picrosirius Red (Polysciences, Warrington, PA) to specifically identify fibrillary collagens I and III. HA was detected with a biotinylated, recombinant, δ-heparin-binding motif, TSG6-Fc (HTI-601) (Halozyme Therapeutics, San Diego, CA) (30). Chondroitin sulfate was identified by immunohistochemistry (Ab11570, Abcam, Cambridge, MA). To quantify specific extracellular matrix components, reacted slides were scanned on a Nanozoomer Digital Pathology slide scanner (Hamamatsu; Bridgewater, New Jersey) and the digital images imported into Visiopharm software (Hoersholm, Denmark) for analysis. Regions of interest were identified using the Image Analysis module and verified manually. The software was programmed to distinguish positive from negative reactivity using a customized algorithm on thresholded pixel values. Images were processed in batch to generate percent areas of positive staining in square microns.

Hyaluronan content

Tissue samples were digested with proteinase K (Sigma-Aldrich, St. Louis, MO) for 24 h at 50–55°C. The enzyme was heat-inactivated and the samples centrifuged at 12,000 rpm for 10 min. HA levels in the supernatant were determined by HA DuoSet ELISA (R&D Systems, Minneapolis, MN) per the manufacturer’s instructions.

Tissue water content

Water content was determined by desiccation (31). Harvested tissues were weighed and dried at 50°C until a constant weight was achieved (typically 48 h). Additional samples were dried for 96 h to confirm that desiccation was complete. A subset of tissues was also incubated at higher temperatures for up to 96 h with comparable results. Tissue water content was calculated as a percentage of the total tissue wet weight from the difference between the wet and dry weights.

PEGPH20 treatment

PEGPH20 was formulated as previously described (28). Pegylation extends the circulatory half-life of recombinant human hyaluronidase (PH20) from <3 min to >10 h (28). Briefly, PH20 was pegylated by conjugating the N-hydroxysuccinimidyl ester of methoxypoly(ethylene glycol)-butanoic acid (i.e., PEG) to PH20 (10:1 molar ratio of PEG/PH20). Purified preparations of PH20 had a specific activity of ∼100,000 U/mg; pegylation (PEGPH20) decreased the specific activity ∼3-fold (31,000 U/mg). PEGPH20 (15 mg/kg) was administered intravenously or intratumorally by bolus injection; control animals received vehicle injections (10 mM histidine and 130 mM NaCl, pH 6.5).

Substrate specificity of enzyme preparations

The substrate specificities of PH20 and PEGPH20 were interrogated with the following GAGs: high-molecular-mass HA (1.2 MDa; HA15M-1, Lifecore Biomedical, Chaska, MN); chondroitin sulfates A and C (400658-1A and 400670-1A, respectively, Amsbio, Cambridge, MA); and chondroitin B (C3788, Sigma-Aldrich). Reactions contained HA (400 μg/mL) or chondroitin sulfates A, B, or C (2 mg/mL). PH20 or PEGPH20 (1,000 U/mL) was incubated with substrate at 37°C for 1.5 h; reaction products were identified by electrophoretic separation on 4–12% TBE gels (Life Technologies, Carlsbad, CA), which were then incubated overnight in 50 μg/mL Stains-all/50% ethanol solution (Sigma-Aldrich) and destained for 2 h in dH2O.

Statistical analyses

Data are expressed as the mean ± SE unless stated otherwise. Unpaired t-tests were performed to compare normally distributed data sets, except for cardiac cessation experiments, which were analyzed by paired t-tests. Multigroup data were analyzed using one-way analysis of variance followed by a Tukey’s multiple-comparisons post-test.

Results

We sought to understand the basis for the unusually high IFPs in PDA (6). To directly investigate the contributions of distinct components of interstitial fluid to these pressures, we performed measurements with both WN and PC in an autochthonous, genetically engineered mouse model (GEMM) of PDA and a variety of other experimental tumor models (Fig. S1 in the Supporting Material).

The GEMM of PDA is based on the targeted endogenous expression of activated KrasG12D and point mutant Trp53R172H to progenitor cells of the developing pancreas (26, 27). The model faithfully recapitulates the clinical manifestations, histopathology, and genetic progression of the human disease from inception to invasion and metastasis (reviewed in (32)). Both primary tumors and metastases in these KPC mice express extremely high levels of interstitial HA (6). We performed IFP measurements in KPC mice possessing large primary tumors ranging between 5 and 9 mm in diameter (for comparison, the normal murine pancreas is ∼15–20 mm in length; see example in Fig. S1, A and B). Consistent with our previous observations (6), we found very high fluid pressures measured with the PC (76 ± 4.2 mmHg, n = 7) (Fig. 1 A). PC-measured pressures in the normal pancreas are 2.6 ± 0.44 mmHg (n = 6) and −0.73 ± 0.6 mmHg (n = 4) in the head and tail of the gland, respectively.

Figure 1.

Figure 1

Assessing fluid pressures in normal tissues, autochthonous PDAs, and transplanted tumor models. (A) The PC measures significantly higher fluid pressures than the WN in autochthonous KPC PDAs. ∗∗∗∗p < 0.0001. (B) The PC also measures significantly higher IFPs than the WN in allografts and xenografts that have increased HA content. Note that approximately equal pressures are seen only in B16-F10 xenografts, which have low HA content. Subcutaneous allografts were made from KPC PDA cells (mPDA#1 and mPDA#2), NIH3T3 cells (3T3), and B16-F10 melanoma cells (B16); xenografts were made from human HT-1080 cells (HT) and BxPC3 cells (BxP). ∗∗∗∗p < 0.0001; ∗∗p < 0.01; p < 0.05.

Parallel measurements performed with the WN in the same autochthonous PDAs revealed much lower pressures (12 ± 3.0 mmHg, n = 5) compared to PC measurements. For comparison, the WN-measured pressure is −0.22 ± 0.33 mmHg (n = 8) in the head and −0.60 ± 0.55 mmHg (n = 4) in the tail of the normal pancreas. The WN-recorded pressures in autochthonous PDA are even lower than the free-fluid pressures previously reported with the WN across a range of subcutaneously engrafted tumors (reviewed in (5)). Free-fluid pressures of this magnitude would not be expected to appreciably influence fluid mechanics in PDA and certainly could not explain the widespread vascular collapse characteristic of these cancers.

Elevated IFPs in transplanted tumors correlate with HA content

We next investigated interstitial pressures across a range of subcutaneously engrafted tumors from both human and murine established cell lines (Fig. 1 B). These experiments also afforded the opportunity to test the relationship between fluid pressures and HA across a wide range of concentrations (Table 1). Several observations were noteworthy. First, autochthonous KPC PDAs have significantly higher IFPs as measured by the PC than allografts of primary cell lines from the same tumors (43 ± 9.0 mmHg and 44 ± 5.3 mmHg, respectively, for the two independently derived cell lines shown; p < 0.05 for comparison to autochthonous pressures). Second, the pressures measured by the WN, which ranged from 4.7 to 14.2 mmHg, were again substantially lower than those measured by the PC. Third, the highest PC-recorded fluid pressures were observed in tumors with the highest HA content; the widest discrepancies in pressures recorded by the two instruments occurred in tumors with high HA content (Fig. 1 B; Fig. S2, A and B; Table 1).

Table 1.

HA Concentration in Tissues and Tumors

Tissue [HA] (ng/mg)a n
Normal pancreas (head) 34 ± 2.7 3
Normal pancreas (body/tail) 23 ± 1.9 3
Lung 7.8 ± 1.4 3
Spleen 4.6 ± 2.1 3
Kidney 1.9 ± 0.2 3
Liver 0.3 ± 0.01 3
KPC PDA 420 ± 150 4
mPDA#1 (allograft) 362 ± 59 6
mPDA#2 (allograft) 169 ± 52 3
NIH3T3 (xenograft) 192 ± 10 3
BxPC3 (xenograft) 267 ± 45 7
HT-1080 (xenograft) 61 ± 4.6 3
B16-F10 (xenograft) 6.3 ± 1.5 3
a

Mean ± SE.

There is sufficient biological heterogeneity across tumor types—and even within independent examples of a given tumor type—to preclude a rigorous mathematical equation to predict IFP from HA concentration. However, a semiquantitative relationship between IFPs and HA concentration can be developed that is consistent with the presumption of at least two components of fluid pressure: a rapidly increasing, saturable component and a more slowly and linearly rising, unsaturable component (Fig. 2, A and B). We propose that the hyperbolic portion of the relationship between IFP and HA concentration is driven largely by the avid binding of water to HA and the swelling pressure exerted by this highly negatively charged polymer; this pressure is constrained—up to a point—by active contractile forces exerted by a tethered collagen-microfibrillary network as cells attempt to maintain tensional homeostasis in response to the applied load. The slowly rising, linear component, by contrast, reflects increases in fluid pressure from the more typical oncotic forces associated with macromolecules, including the Donnan potential. In principle, the pressures measured in the gel-fluid phase should be transmitted uniformly throughout the free-fluid phase in equilibrium with it, but that appears not to be the case here (and see below). Thus, either the WN fails to accurately report the fluid pressure transmitted across these two phases or the PC measures something else.

Figure 2.

Figure 2

IFP as a function of HA concentration. (A) IFP measured by the WN across a number of allografted and xenografted tumors can be fit to a linear relationship as a function of HA concentration. Data points represent independent tumor grafts from the following cell lines (left to right): B16-F10, HT-1080, mPDA#2, NIH3T3, BxPC3, and mPDA#1. (B) IFP measured in parallel by the PC in the same model systems as in (A) demonstrated a hyperbolic relationship between IFP and HA concentration. The data could be well fit according to a saturation isotherm described by IFPPC = IFPmax ([HA]/([HA] + K)).

Measuring hydrostatic pressure in viscous fluids

We sought to further characterize the behavior of the PC and the WN in a series of controlled in vitro settings involving various configurations and concentrations of pure HA in solution and free fluid. We began by adding stepwise increments of a concentrated HA solution to free water in a closed tube: aliquots were carefully added on top, underneath, or directly into the free-water phase in which both instruments were recording simultaneous pressure measurements (Fig. 3). In the first configuration, aliquots of HA were added above the water fluid level without mixing (Fig. 3 A). As can be seen, both instruments tracked the stepwise increases in pressure identically (Fig. 3 B). Next, a barrier of HA was first introduced into the tube and aliquots of water were gently added on top without mixing (Fig. 3 C); the hydrostatic pressures generated and transmitted in this manner are rapidly and completely captured by the PC but not by the WN (Fig. 3 D). Finally, aliquots of concentrated HA were added with vigorous mixing in between; even in the setting of these more homogeneous solutions (Fig. 3 E), the WN struggled to fully capture the fluid pressures in this system comprised of only pure HA and water (Fig. 3 F). Similar results were observed with the nylon threads removed from the WN. Increasing the ionic strength to 2 M NaCl blunted the difference in pressures measured by the PC and WN, whereas chelation with 10 mM EDTA widened it, consistent with shielding versus unmasking, respectively, the electrostatic repulsive charge (not shown).

Figure 3.

Figure 3

Responses of PC and WN in free-fluid solutions of water and pure HA. (A) The PC and WN were inserted into pure water in a sealed vacuum tube to which sequential aliquots (green cross-hatching) of a concentrated 1% HA solution were carefully added without mixing. (B) The PC (blue line) and WN (red line) measured the same pressures with similar kinetics after injections of HA (arrows) as diagrammed in (A). (C) The PC and WN were inserted into a concentrated solution of 1% HA in a sealed vacuum tube. Sequential aliquots of water were then gently layered on top of the HA. (D) The WN (red line) did not fully capture hydrostatic fluid pressures that were transmitted through a gel-fluid phase after injections (arrows) of pure water (diagrammed in C). The PC (blue line) tracked pressures reliably under the same conditions. (E) The PC and WN were inserted into pure water in a sealed vacuum tube, and sequential aliquots of 1% HA were added and then rapidly mixed to generate a homogenous solution. (F) Compared to the PC (blue line), the WN (red line) demonstrated a marked lag in response time and a widening gap in final pressures with increasing HA concentration, despite rapid mixing after each aliquot of HA (arrows).

Measuring hydrostatic pressure in hydrogels

In a second set of experiments, we tested the ability of these instruments to track pressures in sodium polyacrylate, a chemically cross-linked, true hydrogel (Fig. 4 A). Similar to HA, this extended, water-avid polymer has a highly negatively charged backbone and expands from its compacted, dehydrated state upon binding water (Fig. 4 B). Unlike HA, however, it is cross-linked, which constrains its expansion. Given its considerably dehydrated state at the outset, we expected the WN to record negative (subatmospheric) fluid pressures until sufficient free fluid accumulated in continuous channels to register atmospheric pressure. The WN was inserted into a dehydrated cube of sodium polyacrylate and the cube was placed in a container of excess free fluid. The polymer avidly imbibed water and swelled; the WN pressure reached a nadir within ∼2 h, remained there for several hours, and then rapidly rose to atmospheric pressure (Fig. 4 C). This curve also followed the expected changes in compliance of the substance at various stages of hydration: low initial compliance (and therefore marked changes in pressure with changes in volume) followed by a period of extremely high compliance in which volume increases substantially with little to no change in pressure, and, lastly, a low compliance state at maximal hydration accompanied by a significant increase in pressure. A very similar evolution in compliance with increasing hydration is seen in the subcutaneous tissue space (33, 34).

Figure 4.

Figure 4

Responses of PC and WN in a nonuniformly expanding hydrogel. (A) Schematic structures of sodium polyacrylate in dehydrated and hydrated states. Sodium polyacrylate is tightly packed and cross-linked in its dry form. As it absorbs water, the sodium ions disassociate, exposing negatively charged carboxyl groups that form hydrogen bonds with water molecules and cause the polymer to expand. (B) Images of a dehydrated (left), partially hydrated (middle), and fully hydrated (right) cube of sodium polyacrylate after incubation in excess water for 0, 1, and 12 h, respectively. (C) The WN was inserted into a dehydrated cube of sodium polyacrylate and the cube was then placed in a pool of pure water. The WN measured a rapid initial decline in pressure as fluid was imbibed by the cube. This was followed by a slow increase in pressure and then a rapid return to atmospheric pressure. The solid and dashed lines represent two independent experiments. (D) The PC measured both positive and negative pressures under the same conditions as in (C), indicative of nonuniform expansion and binding of water. The solid and dashed lines represent two independent experiments. (E) Schematic representation of the expanding hydrogel tested in (C). As the hydrogel imbibes water and swells, fluid channels begin to form and then coalesce, exposing the probes to free fluid. After several hours, the gels become fully hydrated and reach equilibrium with the surrounding water.

Pressures measured by the PC are influenced by the same processes, albeit less predictably, as different portions of the cube expand at differing rates (Fig. 4, D and E). The initial compacted state gives rise to positive pressures, which then fall and rise as the cube swells nonuniformly. We would predict that simultaneous pressure measurements taken at multiple points throughout this nonhomogeneous gel-fluid state would sum to zero during expansion of the cube and then all read at or near zero in the homogeneous solution at steady state. Indeed, the two experiments shown with random placement of the PC are consistent with this prediction. The final pressures measured by the PC were 2.1 and 1.9 mmHg, respectively.

Substrate specificity of a modified recombinant enzyme

Since its discovery as a “spreading factor” in testes extracts, purified preparations of hyaluronidase have been studied extensively and used to facilitate the subcutaneous absorption of fluids (hypodermoclysis) and substances (35, 36, 37, 38, 39) and also administered intravenously to deplete intracardiac HA and reduce myocardial ischemic injury (40, 41, 42). Although extremely effective in these capacities, the animal-sourced enzyme can induce hypersensitivity reactions and has a very short circulatory half-life. A pegylated recombinant form of the enzyme was developed to overcome these limitations and to extend the potential usefulness of this approach (28).

Before testing the effects of targeted depletion of HA on pressures in the varying tumor models, we first confirmed the specificity of pegylated recombinant hyaluronidase (PEGPH20) for its substrate. In particular, we wished to determine whether other GAGs, primarily chondroitin sulfate (CS), might also be degraded by this intervention and, by implication, represent additional contributors to basal gel-fluid pressures. Although it is well established that hyaluronidase preferentially targets HA over other substrates (43, 44, 45), it can, in principle, degrade other GAGs under certain conditions (46). Both the purified recombinant human enzyme (PH20) and its pegylated form preferentially degraded HA over chondroitin sulfates A, B, and C at pH values between 6.5 and 7.4 (Figs. S3 and S4). Indeed, the enzymes become less discriminating only at pH values well below the physiologic range (i.e., pH ≤6.0), which are not encountered even in the relatively more acidic environments found in carcinomas (47, 48). Available reagents also permit immunohistochemical assessments of CS-A and CS-C in tissues; we found that CS expression was very low at baseline in tumor xenografts and that the small amount present did not change appreciably after PEGPH20 treatment (Fig. S5, A and B). We note that depleting HA did not affect solid tumor components: neither collagen content nor cell density changed acutely after PEGPH20 treatment (Fig. S5 A).

Free fluid and gel fluid in xenografts and autochthonous PDA: targeted depletion of interstitial HA liberates the gel-fluid phase

We have hypothesized that the majority of interstitial fluid in PDA is bound to HA in a less mobile, gel-like state due to entanglement of HA strands, and that the PC—but not the WN—accurately measures pressures transmitted through this state (6, 29, 49). To test these hypotheses, we first measured total tissue-water content in several normal tissues, a range of xenografted and allografted tumors, and in both autochthonous primary tumors and tissues with a high metastatic tumor burden from KPC mice. In all neoplastic tissues, there was significantly increased total tissue water compared to normal controls, as expected (Fig. 5, A and B). We would also expect that in this state, the tissue is sufficiently hydrated for the PC to be minimally influenced by compacted elements (i.e., solid stress).

Figure 5.

Figure 5

Water content in normal tissues, autochthonous PDAs, and subcutaneous allografts and xenografts. (A) Total tissue water is increased in autochthonous PDAs, tumor allografts, and xenografts compared to normal (WT) pancreas. Autochthonous PDA (KPC), as well as all allografts and xenografts, had significantly higher water content than normal pancreas. ∗∗∗∗p < 0.0001. (B) Significantly higher fluid content was measured in the pancreas and liver of KPC mice with primary tumors and metastases compared to the respective normal (WT) organs. ∗∗∗∗p < 0.0001.

We generated subcutaneous tumor grafts from a human PDA cell line (BxPC3) that was transduced to overexpress HA synthase 3 (HAS3) and secrete larger amounts of HA. The xenografts presumably also contain sufficient interstitial free fluid in communication with an abnormally “leaky” intravascular compartment for the WN to function at or near optimum. BxPC3/HAS3 xenografts did indeed have a further increase in water content (85%, n = 7) compared to their nontransduced counterparts, which were themselves fluid-rich (81%, n = 8) (p < 0.0001). We next performed real-time, intravital IFP measurements in these xenografts with the PC and WN before and during systemic administration of PEGPH20 and charted the amplitude and kinetics of pressure changes in the distinct fluid phases. We note first that basal IFP measured by the WN is higher in xenografts than in autochthonous PDA, consistent with more abundant free fluid in the interstitium due to an abnormal, fenestrated (leaky) vasculature characteristic of engrafted tumor cell lines. A small (2–5 mmHg), rapid, and reversible increase in pressure is captured by both instruments immediately after the intravenous bolus injection, consistent with a transient increase and subsequent reequilibration with the hydrostatic (i.e., free) fluid pressure in the microvasculature (Fig. 6 A). Fluid pressures rapidly decrease over the next 20 min followed by a slower decline. At steady state after PEGPH20 treatment, the pressures measured by the two methods equalized, consistent with the gel fluid that was previously in complex with HA being mobilized and liberated after HA degradation. Further, and also consistent with this interpretation, total intratumoral HA (Fig. 6 B; Fig. S5 A) and water content (Fig. 6 C) both decreased significantly after treatment with PEGPH20. Thus, the remaining fluid pressure, comprised now mostly of free fluid, is measured equally well by either instrument (Fig. 6 A).

Figure 6.

Figure 6

Targeted depletion of HA releases water from the gel-fluid phase and lowers fluid pressures. (A) Real-time, intravital IFP in BxPC3/HAS3 xenografts measured by PC (blue line) and WN (red line) before and during systemic treatment with PEGPH20 (arrow). The significantly higher baseline IFP values measured by the PC converge with those measured by the WN after targeted depletion of HA and mobilization of complexed fluid. (B) HA content in BxPC3/HAS3 tumors 2 h posttreatment with vehicle (V) or PEGPH20 (P), demonstrating significant depletion in enzyme-treated tumors. ∗∗∗∗p < 0.0001. (C) Water content of BxPC3/HAS3 tumors 2 h after vehicle or PEGPH20 treatment, showing significant reduction in tumors from enzyme-treated animals. ∗∗∗∗p < 0.0001. (D) Real-time, intravital IFP in autochthonous PDA measured by PC (blue line) and WN (red line) during intravenous treatment with PEGPH20 (arrow). (E) Real-time, intravital IFP in autochthonous PDA measured by PC (blue line) and WN (red line) after intratumoral injection of PEGPH20 (arrow). (F) Cardiac cessation had modest effects on interstitial fluid pressures measured with either the PC (blue symbols) or WN (red symbols). Each line connecting a pair of points represents an independent animal. p < 0.05. (G) The mean change in IFP after cardiac cessation recorded by the PC (blue symbols) in autochthonous KPC PDA and xenografts was −3.5 ± 1.1 mmHg and −2.1 ± 0.5 mmHg, respectively. The WN-recorded pressures (red symbols) changed by an average of −1.3 ± 0.9 mmHg in autochthonous PDA and −1.7 ± 0.9 mmHg in xenografts.

In autochthonous PDA with high interstitial HA concentrations and a functionally and structurally intact vasculature, we hypothesize that free fluid is limiting, hence the very low pressures recorded by the WN. In this case, we expected the WN-measured pressure to instead rise as fluid is mobilized from the bound compartment. We delivered the pegylated hyaluronidase (PEGPH20) by two different routes into the tumor bed: intravenously (Fig. 6 D), and by direct intratumoral injection (Fig. 6 E). Both routes gave the same overall result, albeit with distinct kinetics. Intratumoral injection of PEGPH20 led to a very rapid drop in PC-measured pressure with a corresponding increase in pressure measured by the WN. Systemic PEGPH20 administration also caused a drop in PC-measured pressure with a slightly slower time course, and once again, the WN-measured pressure increased (Fig. 6 D).

Cardiac cessation does not completely collapse IFPs

The above studies were designed to investigate the properties and mechanisms of increased pressures responsible for the hypoperfused, largely collapsed vasculature of PDA and, as a corollary, the precise features measured by the PC and the WN under various conditions. The results of these experiments raise some question about whether the pressures measured by the WN are, in fact, in equilibrium with vascular pressures. To address this directly, we arrested the heart in anesthetized animals and followed the resultant impact on both WN- and PC-recorded interstitial pressures (Fig. 6, F and G). It has been theorized, though not demonstrated, that cardiac arrest should cause pif to go to zero in tumor xenografts (13, 50, 51). We found that by either instrument, although pressures did drop, the changes were small: the WN-measured pressures decreased somewhat more in implanted than in autochthonous tumors, again perhaps suggesting somewhat better communication with the vascular compartment in this setting. In virtually no case, however, did the WN-measured pressure go to zero, consistent with previous reports (13, 50, 51) and despite theories to the contrary. Thus, either the WN-measured pressures are not in equilibration with the vascular space (9) or free-fluid pressures remain positive even after cardiac contractility ceases, a plausible interpretation given that a hydrostatic column of fluid remains in the circulation despite the absence of active pumping: although transvascular fluid flux would be expected to fall to zero under these conditions, particularly as the oncotic pressure gradient is dissipated in the setting of leaky vessels, the capillary and interstitial free-fluid hydrostatic pressures need only equalize, not reach zero. We conclude that total intratumoral (noncellular) water is therefore divided among at least three compartments in equilibrium: intravascular, interstitial free, and interstitial gel. The relative fraction of fluid in each compartment will define the types of interventions most likely to succeed in alleviating prohibitively elevated pressures (Fig. S6).

Discussion

Methods and models

We have shown here that interstitial fluid in autochthonous PDA, as well as many transplantable tumor models, is found predominantly in complex with HA (in both soluble and potentially “entangled” fractions) together with a minor component of free fluid. These investigations also explain why the unexpectedly high levels of gel fluid in the PDA interstitium have until recently gone unnoticed. Prior studies of interstitial fluid, dating back to the seminal work of Starling (1), have focused almost exclusively on the freely mobile component. This focus has persisted until very recently for a number of reasons: 1) all of the early instrumentation and variations thereof, including needle, WN, and perforated capsule among others, could measure only freely mobile fluid; 2) the principal pathophysiologic condition of interest through the midcentury was dropsy, or edema, i.e., the accumulation of excess free fluid in the tissue spaces; and 3) the models of malignancy studied consisted largely of tumor balls of established cancer cell lines grown in 3D-culture systems, or as xenografts or allografts, which have biophysical properties that are very distinct from autochthonous cancers (reviewed in (49)). Solute transport properties have been characterized extensively in various in vitro systems and transplanted and explanted tumor models (52, 53), but they have rarely been investigated in autochthonous cancers (49). It is perhaps not surprising, therefore, that this large body of work has elaborated detailed principles of free-fluid mechanics while almost completely ignoring the predominant, and far more physiologically relevant, HA-dependent gel-fluid phase.

The autochthonously generated cancer neo-organ is only poorly approximated by in vitro and engrafted tumor models. This is particularly true when compared to a complex, stromal-rich cancer such as PDA. Subcutaneously or orthotopically transplanted tumors generated from established invasive cell lines develop rapidly (days to a few weeks), with a correspondingly rapid expansion of a nonnative vasculature. The vessels in such xenografts and allografts possess an increased permeability and hydraulic conductivity from fenestrations and open interendothelial junctions; these, in turn, result in excess free fluid beyond that encountered in normal tissues or autochthonous cancers. Our data suggest that even in these artificial settings of compromised vascular integrity, the majority of the fluid appears to be in a less mobile phase. Autochthonous pancreas cancer vessels have demonstrably few fenestrae and open interendothelial junctions and, as a result, exhibit passive transport properties similar to those of normal pancreas vessels (15). Indeed, the profound differences in physiology and pathobiology between transplanted and autochthonous cancers (54, 55, 56) have direct implications for drug perfusion (14) and corresponding responses to therapy (57). It remains to be seen how many other autochthonous cancers also differ from their transplanted counterparts in this important regard.

Pressures in context

Our studies raise questions regarding what the WN and PC actually measure in a given context. The specific contributions of free- and gel-fluid pressures to total interstitial pressure vary according to the structural and functional integrity of blood vessels, the composition and precise architecture of the extracellular matrix, and the overall cellular content of an organ or tumor; these parameters can differ widely across tissues, tumor types, and model systems. In many normal organs, the interstitial free-fluid pressure is negative and is thought to result from the effects of an unsaturated gel-fluid phase and structural elements in the tissue (25, 33). The negative pressure has been equated to a “tissue absorption” or “imbibition” pressure (16) and can exist only when the interstitium is in a dehydrated state (7, 33). In this state, the structural elements, comprised of the collagen-elastin network, are thought to be compacted (i.e., under compression) and respond by expanding. Given unfettered access to saline, interstitial tissues will imbibe fluid and swell, and interstitial free-fluid pressures will rise to become atmospheric (i.e., zero) as the structural elements now come under tension rather than compression (33). At steady state, therefore, the interstitial free-fluid pressure equals the arithmetic sum of the hydrostatic and oncotic pressures in the gel-fluid phase.

It has been suggested that the PC is subject to solid pressure artifacts (58, 59); the PC measures pressure by displacement of a piezoelectric membrane sensor and so can, in principle, be influenced by solid stress. However, that it can be does not mean that it always is, and whether it measures predominantly fluid pressure or solid stress depends on the particular context and precautions taken in using the instrument. It is possible under conditions of dehydration, such as those found in some normal tissues and in the experiments performed here with sodium polyacrylate, that direct contact with solid elements can occur. The slightly positive pressures measured by the PC in the normal pancreas may conceivably reflect a contribution from solid tissue elements, although they may also arise from gel-fluid pressure (note that the PC-measured pressures in the head versus the tail of the gland also correlated with HA content). In any case, under conditions of normal to overhydration, these elements are no longer compacted, and the PC should no longer be in direct contact with them. We also perform checks before and after to ensure that solid stress does not contaminate the pressure recordings (see detailed descriptions of procedures in (6, 29)). Other investigators have examined the possibility that the PC is affected by solid stress and found that not to be the case (60). The identical pressure readings measured by the PC in tumor xenografts in both sheathed (i.e., “PC-in-needle”) and unsheathed configurations further demonstrate that the instrument is not influenced by solid stress in these settings (29). Thus, we propose that under conditions of an abundant gel-fluid phase that is well hydrated, the PC captures hydrostatic pressure associated with and transmitted through the gel-fluid phase. Under these same conditions, the WN does not accurately report these transmitted pressures.

What exactly, then, do the PC and the WN measure in the distinct natural and artificial contexts described here? The experiments in the simplified system of pure HA in solution suggest an answer that also appears to satisfy the behavior observed in vivo. If we apply the two-phase model of free fluid and gel fluid in the interstitium (10, 33), then at steady state,

pifπifPigΠig,

in which pif and πif refer to the hydrostatic and oncotic pressures, respectively, in the interstitial free-fluid phase, and Pig and Πig refer to the corresponding pressures in the gel-fluid phase. The difference in hydrostatic pressures between the two phases is balanced by the difference in oncotic pressures:

PigpifΠigπif.

In the in vitro experiments in pure HA solutions, there is no free-fluid colloid protein and, assuming that there is little to no HA in the “free fluid” fraction, we can further approximate the difference as

PigpifΠig.

Note that this expression is the same as that used by Brace and Guyton (16) for the tissue absorption pressure (i.e., pif) measured with implanted capsules:

pifPigΠig.

We know that the WN captures pif. The PC, on the other hand, can measure the far higher hydrostatic pressures associated with and transmitted through the gel-fluid phase, i.e., Pig.

Therefore,

PigpifPCWNΠig.

In fact, the difference of 30–40 mm Hg in the hydrostatic pressures measured here at higher HA concentrations is consistent with values reported in the literature for oncotic pressures of soluble HA preparations (16, 61, 62). Applying the more general version of this expression to the in vivo conditions in both autochthonous and transplanted tumors, we have

PigpifPCWNΠigπif.

Now we are left with a question of terminology and of emphasis on the relevant physicochemical properties of the various interstitial phases and processes. How shall we describe the pressures associated with soluble HA? Unlike other GAGs, HA is neither bound nor immobilized in the interstitium (63, 64). Because HA cannot be cross-linked, it does not form a hydrogel (65). Rather, HA in solution behaves as a classical Newtonian fluid over a wide range of concentrations and shear rates—including those encountered in normal and neoplastic physiology—and becomes non-Newtonian only at the extremes (66). Pressures associated with the gel-fluid phase have historically been categorized as solid stress, because it has an elastic modulus. It is more precise, however, to say that this phase is viscoelastic, possessing the force transmission properties of a fluid and the elasticity of a solid. Moreover, most of the relevant properties relating to the observed effects in vitro and the behaviors in vivo appear to be attributable to hydrated HA itself. Thus, it seems reasonable to refer to the gel-fluid pressure as a fluid pressure.

Vectorial versus hydraulic force transmission

Solid stress may also contribute to total tissue pressure, of course, particularly in the setting of the hypercellular tumors more commonly observed with transplanted cancer cell lines (67). PDAs are notoriously hypocellular, however, especially with respect to the epithelial tumor cell content (68), and increased cell density therefore probably does not explain vascular collapse. Other fixed tissue elements such as collagen did not change appreciably during the relatively short duration of our experiments (i.e., minutes to hours) and so were not the cause of the observed drop in fluid pressures with enzymatic depletion of HA. The large array of negative charges along the HA polymer creates an electrostatic repulsion and a corresponding expansive or swelling force. Indeed, it is important to bear in mind that in addition to resisting compression, hydrated HA also swells (16, 22). This expansion would apply a tensile load to collagen fibrils tethered to stromal and tumor epithelial cells, and these fibrils would, in response, be expected to contract in an attempt to maintain tensional homeostasis (69, 70). Thus, active forces may cooperate with the passive forces described above: solid stress may augment gel-fluid pressures through ATP-dependent contraction of collagen fibrils (Fig. S6). Interstitial pressures would rise less precipitously if hydrated HA was permitted to freely expand. However, even though the expanding HA-dependent fluid phase is constrained by these solid tissue elements, this does not make the source a solid stress; it is still hydraulic. We interpret the hyperbolic relationship between HA concentration and fluid pressure as reflecting this balance between HA swelling and collagen fibril contraction, the latter of which has a limit (Fig. 2 B; Fig. S6). We hypothesize, therefore, that specifically targeting these contractile forces would cause a stepwise decrement in pressure that corresponds largely to the applied electrostatic repulsive force driving HA expansion. Removal of macromolecular HA would be required to further alleviate pressures associated with the more traditional oncotic forces and the Donnan potential (i.e., the unequal distribution of charge across distinct fluid compartments) together with a minor component from Van’t Hoff forces.

It is essential to note that solid pressures are, by definition, limited to the application of point forces (i.e., they are transmitted vectorially at sites of direct contact). With measured pressures on the order of 60–80 mmHg resulting in collapse of ∼75% of vessels, the postulated total force summed over the surface area of the collapsed vasculature would be astronomical and implausible. By Pascal’s law, on the other hand, the pressure applied to a confined fluid is transmitted equally throughout every portion of the fluid (i.e., hydraulic force distribution). We propose therefore that the defining characteristic of this gel phase is not its properties as a “solid,” but its fluid properties or, perhaps better, its viscoelastic properties. It is only by invoking the hydraulic transmission of forces that a thermodynamically plausible and biophysically tenable explanation can be proposed for the observed vascular collapse in PDA. It is for these reasons that we have chosen to refer to the pressure associated with this less mobile fluid—historically also referred to as a gel fluid—according to a functional, rather than a purely categorical, definition and, therefore, as a fluid pressure and not a solid stress, because this property embodies the most biologically and pathophysiologically salient feature.

Clinical implications

It may seem counterintuitive that a macromolecular therapy can permeate sufficiently to degrade HA in the tumor interstitium, particularly as the vessels in these cancers appear to resemble normal vessels both structurally and functionally. However, several points are germane in this regard. First, we note that most capillary beds are characterized by two broad classes of pores: the more abundant small pores, and the so-called large pores, which represent only a few percent of the total. As a result, many large molecules can enter the interstitium: its reflection coefficient of 0.85–0.95 notwithstanding, the albumin in the circulation cycles through the interstitium and is returned via the lymphatics approximately every 24 h (71). Second, we note that purified hyaluronidase has historically been delivered through the circulation and shown to effectively remove interstitial HA (40, 41). The recombinant, covalently modified form used in our studies is minimally pegylated and therefore only a few fold larger than the native enzyme. Third, we note that the systemically delivered PEGPH20 used here also efficiently ablates HA in the interstitium of normal organs and, so, abnormal vessels are not a prerequisite. Indeed, this is true of many macromolecular chemotherapies and contributes to the often considerable unwanted effects that constitute major challenges in clinical oncology. Fourth, although PDAs are hypovascular, they are not avascular: ∼25–30% of the measureable tumor vasculature is patent at baseline, providing pockets of access—and therefore perfusion—within the heterogeneous microenvironment of these cancers (6, 14). Fifth, PEGPH20 was designed to achieve a prolonged half-life (>10 h) in the circulation, permitting extended exposure of the tumor to drug (28). Finally, the agent is an enzyme and therefore acts catalytically, not stoichiometrically, extending its effectiveness even at relatively low intratumoral concentrations. Collectively, these factors permit sufficient enzyme to enter the tumor and catalyze a self-reinforcing process of HA degradation, fluid mobilization, decreased pressure, and vascular reexpansion, followed by increased local perfusion, which then permits further drug entry and so on. PEGPH20 can also induce fenestrae in the tumor vessels it accesses, at least in a GEMM of PDA, further enabling perfusion (15).

This enzymatic strategy appears to be effective in overcoming treatment resistance in human PDA (72). Two national, randomized Phase 2 trials (NCT01839487 and NCT01959139) are presently underway, each testing one of the current standard-of-care chemotherapy regimens for Stage IV (metastatic) PDA in the presence or absence of PEGPH20. We have recently reported an interim analysis of one of these trials revealing an unprecedented objective response rate of 52% in patients whose tumors had high HA content and who received the enzyme, compared to 24% in similar patients who did not receive PEGPH20 (p < 0.038) (73, 74). A significant increase in progression-free survival (9.2 months vs. 4.3 months; p < 0.05) was also observed. Moreover, in patients with low intratumoral HA cancers, the addition of enzyme did not improve chemotherapy efficacy. Finally, among patients who did not receive PEGPH20, those with low HA seemed to fare better than those with high HA tumors, as expected if, indeed, HA impairs perfusion. Each of these pairwise analyses supports the notion that HA and the associated gel-fluid phase are first-line mechanisms of drug resistance in PDA and that specific degradation of this target alleviates a barrier to treatment. A global, randomized Phase 3 study of patients with high intratumoral HA levels has recently begun enrollment (NCT02715804).

Conclusions

We have demonstrated here that IFPs, which can rise inordinately in autochthonous cancers, derive from a small, freely mobile-fluid phase and a dominant gel-fluid phase. In cancers such as PDA, with especially high HA content, pressures associated with the gel-fluid phase cause vascular collapse and hypoperfusion, profoundly altering intratumoral physiology and contributing to therapeutic resistance. Enzymatic degradation of HA can liberate the bound fluid and significantly decrease fluid pressures to levels associated with residual free fluid, thereby permitting vascular reexpansion. Given the potential importance of interstitial gel fluid to cancer pathogenesis and resistance, its effects must be fully accounted for to construct a complete depiction of pressures generated by all phases of interstitial fluid. This understanding can, in turn, rationally inform the development of treatment strategies most likely to succeed while helping to avoid those destined to fail.

Author Contributions

C.C.D. and K.E.D. designed and performed research, analyzed data, and wrote the manuscript; M.A.C. designed and performed research and analyzed data; R.J.O., C.Z., Z.H., R.J.C., and J.S.B. performed research; C.B.T. and C.D.T. designed research and analyzed data; P.P.P. analyzed data; G.I.F. and H.M.S. designed research; and S.R.H. designed and supervised research, analyzed data, and wrote the manuscript.

Acknowledgments

We thank Ashley Dotson for assistance with animal husbandry and care; Elizabeth Van Volkenburgh, Melissa Lacey, and Ingunn Stromnes for assistance with experiments; Martin Whittle for comments on the manuscript; and John D. Potter for helpful discussions and comments on the manuscript. We are indebted to the anonymous reviewers for their excellent suggestions. We thank Shelley Thorsen and Nathan Lee for expert assistance with figure and manuscript preparation. R.J.O., C.Z., Z.H., C.B.T., R.J.C., C.D.T, H.M.S. and G.I.F. are, or were, employees, consultants, and/or shareholders of Halozyme Therapeutics. The content of this article is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

This work was supported by National Institutes of Health National Cancer Institute grants (CA161112 to S.R.H. and CA152249 to S.R.H and P.P.P.), Lustgarten Foundation (S.R.H.), and the Giles W. and Elise G. Mead Foundation (S.R.H.).

Editor: Leslie Loew.

Footnotes

Paolo P. Provenzano’s present address is Department of Biomedical Engineering and Masonic Cancer Center, University of Minnesota, Minneapolis, Minnesota.

Christopher C. DuFort and Kathleen E. DelGiorno contributed equally to this work.

Supporting Material

Document S1. Figs. S1–S6
mmc1.pdf (1.2MB, pdf)
Document S2. Article plus Supporting Material
mmc2.pdf (2.2MB, pdf)

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Supplementary Materials

Document S1. Figs. S1–S6
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Document S2. Article plus Supporting Material
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