| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
Biophys J, September 2002, p. 1650-1660, Vol. 83, No. 3
*E. L. Steele Laboratory for Tumor Biology, Department
of Radiation Oncology, Massachusetts General Hospital and Harvard
Medical School, Boston, Massachusetts 02114; and
Biological Engineering Division, Massachusetts
Institute of Technology, Cambridge, Massachusetts 02139 USA
 |
ABSTRACT |
|---|
 TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
Diffusion coefficients of tracer molecules in collagen type I gels prepared from 0-4.5% w/v solutions were measured by fluorescence recovery after photobleaching. When adjusted to account for in vivo tortuosity, diffusion coefficients in gels matched previous measurements in four human tumor xenografts with equivalent collagen concentrations. In contrast, hyaluronan solutions hindered diffusion to a lesser extent when prepared at concentrations equivalent to those reported in these tumors. Collagen permeability, determined from flow through gels under hydrostatic pressure, was compared with predictions obtained from application of the Brinkman effective medium model to diffusion data. Permeability predictions matched experimental results at low concentrations, but underestimated measured values at high concentrations. Permeability measurements in gels did not match previous measurements in tumors. Visualization of gels by transmission electron microscopy and light microscopy revealed networks of long collagen fibers at lower concentrations along with shorter fibers at high concentrations. Negligible assembly was detected in collagen solutions pregelation. However, diffusion was similarly hindered in pre and postgelation samples. Comparison of diffusion and convection data in these gels and tumors suggests that collagen may obstruct diffusion more than convection in tumors. These findings have significant implications for drug delivery in tumors and for tissue engineering applications.
| |
INTRODUCTION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
Optimal therapy of tumors requires delivery of
sufficient amounts of therapeutic agents to the target cancer cells.
Thus, the agent must penetrate the tumor interstitial matrix (IM), a complex assembly of collagen, glycosaminoglycans, and proteoglycans (Alberts et al., 1994
). Convection through the tumor IM
is poor due to interstitial hypertension, leaving diffusion as the
major mode of drug transport. As anti-cancer therapy focuses
increasingly on larger therapeutics such as liposomes, which are
typically at least 90 nm in diameter (Gabizon et al.,
1998; Kulkarni et al., 1995), and gene
vectors, which range in diameter from 20 to 300 nm (Costantini
et al., 2000), diffusion within the tumor IM becomes a greater
barrier to delivery (Boucher et al., 1998; Jain,
1999; Netti et al., 1999).
Glycosaminoglycans (GAGs), and particularly hyaluronan (HA), are
believed to play a primary but not exclusive role in regulating fluid
movement in the IM (Gribbon et al., 1998; Levick,
1987). However, diffusion of large molecules in tumors has been
correlated to collagen content and organization, but not to HA content
(Netti et al., 2000; Pluen et al., 2001).
These in vivo studies correlated matrix composition to diffusive
hindrance, but the biological complexity prohibited detailed analysis
of the mechanisms of transport hindrance within the tumor IM. For
example, even within a given tumor, Pluen et al. (2001)
found varying degrees of collagen organization and heterogeneous
distribution of different matrix molecules.
To overcome these problems, we measured diffusion and hydraulic
conductivity in pure collagen type I gels and compared these results
directly with previously published results for tumors of comparable
collagen concentration. Furthermore, we compared the structure of the
gels with that seen in tumors. To investigate the role of collagen
structure, we compared diffusion in collagen gels and solutions of the
same concentrations. The findings presented here are important to the
development of improved drug delivery strategies (Jain, 1998) and to
pharmaceutical applications of collagen matrices, including the design
of tissue substitutes and controlled release devices (Sano et
al., 1998).
| |
MATERIALS AND METHODS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
Experimental techniques
Preparation of collagen gels
Vitrogen 100 collagen type I solution was purchased from Collagen Corp. (Cohesion Technologies, Palo Alto, CA) at a concentration of
3 mg/ml. The pH and ionic strength were adjusted by
addition of NaOH (pH 7.4) and 10× phosphate buffered saline (PBS). To
concentrate the solution, the collagen was ultracentrifuged (Beckman
LC-300) at 10°C for 26-48 h for preparation of 10-45 mg/ml gels.
Supernatant was extracted and pellets were maintained at 4°C.
Collagen concentration in the pellet was determined from the difference
between precentrifugation and supernatant collagen content as
determined by UV spectrophotometry. Pellet concentration was adjusted
by dilution with PBS. The polymerization of highly concentrated
collagen solutions leads to the formation of fibers and filaments. To
obtain a collagen gel formed predominantly of fibers, 30 ml of
neutralized collagen type I (0.4 mg/ml) was polymerized at 32°C for
48 h. The collagen was centrifuged at 11,000 or 25,000 RPM for 12 or 30 min, respectively. The collagen gel was collected on a plastic
coverslip that was attached to the bottom of the centrifuge tube. To
determine the organization of the fibers and the dimensions of the gel,
second harmonic images of the collagen were obtained with a multiphoton
microscope (Williams et al., 2001). The collagen
concentration estimates were based on the unpolymerized collagen volume
and the final gel volume after centrifugation.
For fluorescence recovery after photobleaching (FRAP) experiments at
low collagen concentration (2.4 mg/ml), capillary tubes were partially
filled with collagen solution and kept for 2 h in a 37°C
incubator. After gelation, an aqueous solution of tracer molecules (2 mg/ml) was added to the capillary, which was then sealed and maintained
overnight at 37°C to allow tracer penetration of the gel. For FRAP
experiments with more concentrated gels, the appropriate tracer
molecule solution was added during adjustment of the pellet
concentration. The samples were then prepared on concave microscope
slides under coverslips and sealed with silicone grease. Samples for
permeability and visualization experiments were prepared in Transwell
(24 mm diameter; for 0.24% gels) or Snapwell (12 mm diameter, for 1%
gels) membrane-bottomed cell culture chambers (Corning Costar Corp.,
Cambridge, MA) and maintained in a 37°C incubator for at least 1 h to allow gelation. PBS was then added to chambers to maintain hydration.
Preparation of hyaluronan solutions
Hyaluronic acid sodium salt isolated from rooster comb (Sigma Chemical Co., St. Louis, MO) was dissolved by slow addition of 1× PBS (pH 7.4) for a final concentration of 4 mg/ml. Fluorescent markers at a concentration of 2 mg/ml were added to the solution. The solution was stirred at 4°C for 10 h and subsequently stored at 4°C overnight. Samples were prepared and sealed in capillary tubes as described above for low-concentration collagen gels.Measurement of diffusion coefficients
Diffusion coefficients were measured using the FRAP with spatial Fourier analysis technique described previously (Berk et al., 1993, 1997). Briefly,
samples permeated with FITC-conjugated tracer molecules were placed on
a microscope stage. Each sample was subjected to brief localized 488 nm
irradiation from a krypton-argon laser, resulting in bleaching of
fluorescence in the irradiated spot (radius ~20 µm). Images were
recorded by CCD camera as the bleached spot recovered fluorescence. The
diffusion coefficient was extracted from the exponential time decay of
the spatial Fourier transform of fluorescence intensity. The diffusion
coefficient for a given sample represents the average of 5-10 FRAP
measurements in the sample. When not specified otherwise, three gel
samples were used to determine the diffusion coefficient of each
molecule-gel combination. Tracer molecules including lactalbumin (LA),
bovine serum albumin (BSA), and dextrans of molecular weights 4.4 K-2 M
were purchased in FITC-labeled form (Sigma). Nonspecific IgG was
purchased unlabeled (Sigma) and subsequently conjugated to FITC using
the Fluo-EX labeling kit (Molecular Probes, Portland, OR).
Collagen gel samples were prepared as described above and maintained at
37°C throughout diffusion measurements. For measurements in
unassembled collagen solutions, samples were maintained at temperatures
between 12 and 17°C by supporting the sample on a metal plate in
contact with an ice pack. Gelation did not occur at these temperatures,
as detected by a lack of OD450 absorbance, indicating no turbidity or
light-scattering in these samples. For both gels and solutions,
temperature was continuously monitored using a thermocouple and
maintained within ± 1°C for all measurements on a given sample.
Measurements were also made in solutions of HA (Sigma) at pH 7.4 and
37°C.
Measurement of Darcy permeability
Permeability was measured by monitoring flow rate through collagen gels under hydrostatic pressure in an apparatus described previously (Chang et al., 2000). Briefly, Transwell or
Snapwell cell-culture chambers containing gel samples supported on a
highly porous membrane were fit snugly into a sample holder and
maintained at 37°C. By adjusting the height of the downstream
reservoir, a constant hydrostatic pressure was applied to force flow
through the gels. The flow was directed through a thin capillary into which one small air-bubble had been injected. Air-bubble motion was
visually undetectable due to the low flow rates through the samples.
Thus, the linear velocity of the air-bubble was monitored by a
photodiode attached to a servo-null motor, which tracked the bubble for
30 min-1 h and was used to determine volumetric flow rates.
Hydrostatic pressure (
P) of 5-15 cm H2O
(depending on sample concentration) were imposed to create flow that
resulted in the lowest measurable bubble velocity. Low concentration
(0.24%) gels were not tested, as they were not sufficiently
viscous/solid. Gels at 1% were cast in Transwell chambers that fit
directly into the apparatus sample holder. Higher concentration gels
(
1%) were cast in Snapwell inserts, and a silicone ring was used to
seal the space between the insert and the outer Transwell support. All
junctions between plastic and gels (collagen or silicone) were sealed
with Krazy Glue to prevent leakage. Leaky samples were quickly detected
due to immediate, rapid movement of the air-bubble and were discarded.
The surface area (A) and thickness (L) of each
sample were measured. The Darcy permeability (K) of the
sample was then determined from the time-averaged volumetric flow rate
(Q) and viscosity (µ) using Darcy's Law:
|
P = 10 cm H2O matched the values
obtained by extrapolating agarose permeability data of Johnson
and Deen (1996) to zero pressure drop (data not shown). To
determine whether the hydrostatic pressure used in these experiments
actually compacted the gels and hence produced erroneous results,
permeability was measured at two different pressures (10 cm and 5 cm
H2O). The ratio of the two flow rates was 2.37±0.86
(N = 12), approximately equal to the expected value of
2, suggesting that compaction was not significant.
Visualization by laser scanning microscopy using either confocal reflectance or second harmonic generation
Samples were prepared as described in Transwell inserts and sealed under a coverslip. Confocal reflectance microscopy was performed using a modified Bio-Rad MRC600 (Bio-Rad Laboratories, Hercules, CA), an Olympus 100× 1.4 NA objective (Olympus America Inc., Melville, NY), and 488 nm light from a Kr-Ar laser (American Laser Corp., Salt Lake City, UT). Reflected light from the back surfaces of the objective was attenuated using a quarter wave plate and an analyzer at the detector (Cheng and Summers, 1990; Friedl et al.,
1997; Brightman et al., 2001). Gels were also
imaged using second harmonic generation (Williams et al.,
2001); 810 nm laser light from a mode-locked Ti:Sapphire was
scanned through a sample using a modified Bio-Rad MRC600, and second
harmonic light was collected using a 405DF33 bandpass filter and an
HC125-02 photomultiplier tube (Hammamatsu, Bridgewater, NJ).
Theoretical models
Effective medium model
To account for hydrodynamic interactions and relate the permeability of a matrix to its diffusive hindrance, Phillips et al. (1989) proposed the Brinkman (or effective medium) model
for a stationary sphere in imposed flow. This model was later modified slightly to account for hindered diffusion in a medium of interest (Solomentsev and Anderson, 1996; Phillips,
2000):
|
is a constant of proportionality
introduced to improve the quality of curve fits to this equation. The
effective medium model, when used in combination with the Carman-Kozeny
model (Carman, 1937) below, was found by Pluen et
al. (1999) to give the best correlation with pore size in
agarose gel experiments.
Carman-Kozeny model
We estimated pore size in gels using the Carman-Kozeny model to relate permeability, K, and pore size, a, for a gel of porosity
:
|
|
|
|
,
of collagen by the equation = 1 
, where is the
product of the collagen concentration and the effective specific volume of collagen (protein + bound water), previously reported as 1.89 ml/g (Levick, 1987).
| |
RESULTS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
Visualization of collagen gels revealed varying degrees of three-dimensional fibrillar assembly
The organization of gels was visualized using a laser-scanning
microscope (in confocal reflectance or 2HG mode). Confocal reflectance
microscopy and second harmonic generation are both performed in
unfixed, hydrated samples, and are useful techniques for the
visualization of the collagen network with a spatial resolution of
~0.5 µm, including distribution and bulk organization of fibers (Friedl et al., 1997; Williams et al.,
2001). No structure was detected in collagen solutions at
12-17°C (data not shown). Fig. 1 shows
the isotropic, three-dimensional nature of collagen gels of
concentrations 0.24% and 4.5%. After gelation, low-concentration gels
(0.24%, Fig. 1 a) show a highly fibrillar organization as seen previously in gels of comparable concentration (Friedl et al., 1997; Brightman et al., 2001). Unlike the
long fibers oriented primarily in two dimensions seen by Friedl
et al. (1997), our gels show more 3-dimensionally oriented
fibers. At higher collagen concentrations studied (1, 3, 4.5%, Fig. 1
b), CLSM revealed poorly organized collagen with denser
arrays of shorter fibers replacing the long fibers seen at lower
concentrations. Inhomogeneous organization of collagen gels prepared
from high-concentration solutions was also seen by transmission
electron microscopy (data not shown) as dense, short-banded structures
alongside unbanded filamentous structures. These observations agree
with previous reports that at concentrations higher than 0.5%,
collagen gels in vitro are formed of a mixture of banded fibrils and
filamentous structures (Williams et al., 1978). All
these gels had an apparent pore size roughly equal to or greater than
the ~0.5 µm spatial resolution of the microscope.
|
When low-concentration collagen solutions were gelated and then centrifuged to a higher concentration, a dense mat of highly fibrillar collagen was formed (Fig. 1 c) with many long fibers compressed close together, with an interfibrillar spacing close to or smaller than the ~0.5 µm resolution of the microscope. Note that the presence of organized structures does not preclude the existence of unpolymerized collagen in what appear to be void spaces.
Collagen gels significantly hinder molecular diffusion
Diffusion data obtained in collagen gels prepared from solutions
of various concentrations are shown in Fig. 2
a, along with data for
diffusion in saline and in HA solution. Results of a one-sample
t-test on slopes of diffusion coefficient versus collagen concentration for representative tracer molecules (dextran 4K, BSA,
dextran 2M) verified that the diffusion coefficients decrease significantly (p < 0.05) with increasing collagen
content. The hydrodynamic radius, Rh, of each
molecule was determined from its diffusion coefficient in solution,
D0, and the Stokes-Einstein relation, under the
assumption that the molecule assumes a spherical configuration:
|
23 J/deg;
T is temperature in K, and µ is the viscosity of water.
|
For reference, correction to 37°C of the diffusion data of
Shenoy and Rosenblatt (1995) in 30 mg/ml succinylated
collagen solution yields comparable results with
D37°C = 2.2 × 107
cm2/s for BSA (Rh = 4 nm), and
D37°C = 2.0 × 107/s
cm2/s for 69 kD dextran (Rh = 6 nm). The linearity of the data sets indicates that the different
classes of tracer particles (globular proteins, dextrans, liposomes)
behave similarly in our experiments, so that particle conformation and
interaction with the matrix do not introduce experimental confounds. In
Fig. 2 b we plot the ratio of the diffusion coefficients
obtained in gels to those in free solution as a function of the
experimental hydrodynamic radius, to more clearly illustrate the
hindrance presented by the gels. The data clearly indicate that at
physiologically relevant concentrations (1-4.5%), collagen poses a
significant barrier to diffusive transport. HA solutions at 0.05% (0.5 mg/ml) showed statistically significantly less diffusive hindrance
relative to the >1% collagen physiological gels studied here
(p < 0.001 for BSA). This HA concentration used was
chosen to correspond to the HA content of the four tumors under
consideration (see below). At much higher HA concentrations (0.4%), we
found significant diffusive hindrance (D/D0
~0.56±0.11 for IgG, D/D0 ~0.27±0.04 for 2M
MW Dextran), equivalent to that found in previous studies (De
Smedt et al., 1994) (data not shown).
Diffusion data in gels closely match previous measurements in tumors
We studied gels prepared from 1% (10 mg/ml), 3% (30 mg/ml), and
4.5% (45 mg/ml) solutions specifically to allow comparison with
diffusion data obtained by Netti et al. (2000) and
Pluen et al. (2001) in the following tumors implanted in
mouse dorsal chambers: human colon adenocarcinoma LS174T, mammary
carcinoma MCaIV, human soft tissue sarcoma HSTS-26T, and human
glioblastoma U87. Measurements by Netti et al. of collagen and HA
content in tumors are given in Table 1.
IM concentrations in these tumors are estimated by approximating the
interstitial volume fraction of the tumor as
fv = 0.20 (Jain, 1987) and
assuming that (1) matrix components are distributed throughout the
interstitial volume, and (2) tissue density is ~1 g/ml. Although the
interstitial volume fraction will vary between tumors, reaching up to
50% (unpublished data) and matrix component distribution is not
uniform within a given tumor, these approximations provide a
rough basis for comparison.
|
To compare diffusion in gels and tumors, we also account for the
tortuosity of the interstitial space resulting from cellular obstacles,
as illustrated in Fig. 3. Diffusion along
an interstitial path with tortuosity 
is reduced according to
DIM = Dgel/2 (Nicholson and
Phillips, 1981; Nicholson and Sykova, 1998).
Tortuosity is difficult to measure and exhibits inter and
intratumor variation. In the absence of detailed data on the
tortuosity of the tumor types in question, the tortuosity of a
well-packed system of cells can be estimated theoretically, although
such a theoretical estimate is a possible source of error. Analytical
and numerical calculations have yielded the value = 21/2 for two-dimensional diffusion in arrays of cells with
negligible intercellular spacing, and for diffusion in a
two-dimensional isotropic pore network (Blum et al.,
1989; Chen and Nicholson, 2000). We use this
value to adjust gel data for comparison with tumor tissue data, because
the FRAP technique measures two-dimensional radial diffusion.
|
In Fig. 4 a-c, we
compare the adjusted gel diffusion coefficients to the data of
Pluen et al. (2001) in tumors of comparable collagen
content. Overall, the gel and tumor data match well, especially
considering the absence of other matrix components in the gel and the
likely differences in collagen organization and distribution between
tumors and gels. The absence of other matrix components may explain the
faster decrease of D with Rh in
tumors than in gels. The difference in slopes is reflected in Fig.
4d, which shows an increase in the effective tortuosity, 
*, is the value of the
tortuosity necessary to completely account for the difference between
the gel and tumor diffusion coefficients, and reflects effects beyond
the geometric considerations discussed above.
|
Gelation of a collagen solution does not significantly affect its diffusional hindrance
Diffusion coefficients were measured in collagen samples pre and postgelation. Measurements were obtained pregelation at 12-17°C and corrected to 37°C using the Stokes-Einstein equation. Confocal reflectance images of collagen solutions verified a lack of observable structure in pregelation samples (figure not shown), which was further confirmed by optical density measurements, which were equivalent to those obtained in water. Pre and postgelation diffusion coefficients were determined for collagen concentrations of 0-4.5%, from multiple measurements within the same sample pre and postgelation, and are shown in Fig. 5. No significant difference was detected between diffusion coefficients pre and postgelation at any of the concentrations of collagen studied, after correction for temperature and viscosity using the Stokes-Einstein relation.
|
Diffusion in gels prepared by centrifuging low-concentration gels does not match diffusion in gels prepared directly from high-concentration solutions
The diffusion coefficient of 2M MW dextran was measured in ~20%
collagen gels prepared by centrifuging previously polymerized 0.04%
gels. The measured value of 4.72 ± 1.7 × 108
cm2 s1 was significantly faster (p < 0.01) than the value of 7.8 ± 5.3 × 109
cm2 s1 measured in collagen gels prepared by
direct gelation of 4.5% collagen solutions as discussed above.
The effective medium model underpredicts the permeability of collagen gels
The Darcy permeability of 1%, 3%, and 4.5% collagen gels was determined experimentally and also estimated from diffusion data using the effective medium model. Curve-fits of the diffusion data to the model are shown in Fig. 2 b, and the experimental measurements and model estimates of the Darcy permeability are compared in Fig. 6 a. The experimental values and model estimates agree only for the 1% gels. Above this concentration, the experimental measurements are increasingly greater than the model estimates, with an order of magnitude difference for the 4.5% gels. This difference in permeability values translates into a difference in pore size as estimated by the Carman-Kozeny model, as shown in Fig. 6 b.
|
Measured permeability of gels does not correspond to tumor permeability
The permeability of gels correlated inversely with collagen
content, whereas the permeability of tumors with corresponding collagen
content did not (Fig. 7 a). To
compare the permeability measurements in collagen gels with the
published measurements in tumors, the gel measurements must be adjusted
by the area fraction (fA) in a tumor slice and
the tortuosity, or increased length of the fluid path through the
slice. Adjusting the gel data by Ktumor = Kgel fA/, where the
interstitial area fraction is estimated at
fA = 0.2 and the theoretical estimate
= 21/2 is used for the tortuosity, we obtained the
data shown in Fig. 7 b alongside published tumor
measurements. Collagen gels corrected for the absence of cells are
significantly less permeable than tumors of comparable collagen
content, although direct comparison may be complicated by the use of
different permeability measurement techniques for gels and tumors, and
by intratumoral ECM heterogeneity.
|
| |
DISCUSSION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
Collagen can account for most of the diffusional hindrance measured in tumors studied
Collagen significantly impedes diffusion, and the extent to which
it does so, when corrected for the tortuosity of the interstitium, is
consistent with diffusion data obtained in tumors of comparable collagen content (Fig. 4 a-c). Note that the
slope of the diffusion data differs between gel and tumor data sets.
This phenomenon is also seen as an increase with molecular size of the
effective tortuosity in tissue (Fig. 4 d), and has been
observed in studies of diffusion in the brain (Nicholson and
Sykova, 1998). In tumors, matrix components other than collagen
could affect this slope by differentially affecting the diffusion of
small versus large molecules. Heterogeneity of collagen structure and
distribution in tumors, as shown by Pluen et al. (2001)
may also differentially affect particles of different sizes. Thus the
effective tortuosity in a tumor scales with particle size and is
heterogeneous, depending on the local tissue composition and structure.
Our results suggest that diffusion in pure collagen gels mimics that in
the tumor IM over the wide range of particle sizes studied. However,
extrapolating these results to particles with a hydrodynamic radius
larger than 2M MW dextrans may not be justified. The Carman-Kozeny
estimates of pore size and the linearity of the diffusion data sets
suggest that the particles we used are smaller than the effective pore
sizes of the gels studied. As particle sizes approach the effective
pore size of the media, the fine structure of the matrix is expected to
critically influence transport hindrance, and in vitro gels may no
longer capture the in vivo behavior. Rusakov and Kullmann
(1998) argued that large molecules comparable to the pore size
experience greater hindrance due to viscous interactions unaccounted
for in tortuosity corrections. Matrix pore size is expected to be
different in gels than in tumors, where factors such as compaction of
collagen fibrils by fibroblasts (Friedl et al., 1997;
Guidry and Grinnell, 1987; Huang-Lee et al.,
1994) and additional IM molecules such as decorin (Pins
et al., 1997) play a role. Thus, although the agreement between
the gel and tumor measurements is surprisingly good, these results should not be extrapolated to larger particle sizes.
Unassembled collagen is implicated in the diffusive hindrance of pure collagen gels
After correction for the effect of temperature on viscosity and
molecular motion, there was no significant difference in diffusion between collagen solutions and collagen gels gelated from equivalent concentrations. These data are consistent with those of Shenoy and Rosenblatt (1995), where solutions of succinylated collagen at room temperature were capable of significantly slowing diffusion. This fact, combined with imaged pore sizes that appear too large (several hundreds of nanometers) to significantly hinder diffusion, suggests that unassembled collagen in the void spaces of these gels
plays a role in hindering diffusion.
Note that gels formed by gelation of different concentrations of
collagen are not simply more or less concentrated versions of the same
structure. The highly fibrillar network formed from the gelation of
low-concentration collagen solutions is qualitatively different from
the dense array of short fibers and partially formed structures
generated upon gelation of high-concentration collagen solutions. When
gels formed from low-concentration collagen solutions (~0.04%) are
subsequently centrifuged to higher concentrations (~20%) than the
gels formed by direct gelation of high-concentration solutions
(~4.5%), the resultant gel retains its original highly fibrillar
structure, but the long fibers are significantly compacted, forming a
dense mat. Not surprisingly, these qualitatively different gels
prepared by centrifugation postgelation do not reproduce the diffusive
hindrance of gels prepared by simple gelation, exhibiting a
significantly higher diffusion coefficient for 2M MW dextran. The
compaction of the array of long fibers initially formed at low
concentrations could be markedly inferior to that of the dense array of
short fibers and partially formed structures generated by gelating a
high-concentration solution. Additionally, it is known that the
partitioning of collagen between assembled and unassembled states
varies with the concentration at which the gel is polymerized
(Williams et al., 1978). We conclude that the poorly
assembled gels formed by simple polymerization of collagen solutions
and containing that proportion of unassembled collagen dictated by the
concentration at time of gelation are the gels that quantitatively
mimic the diffusive hindrance of tumor interstitium of equivalent
collagen concentration.
Although these gels quantitatively mimic the diffusive hindrance of the
tumor interstitium, this does not mean that these gels completely
reproduce the interstitial matrix at a molecular level. Other matrix
molecules are certainly present in vivo, and the structure of collagen
assembled in vivo is likely to differ from that assembled in vitro.
However, the poorly assembled gels studied here do have structural
similarities to the collagen of the tumor interstitium, which is poorly
organized in comparison to normal tissue. Pluen et al.
(2001) reported that subcutaneous U87 tumors stain positively
for collagen type I in the tumor center where only few fibrils were
detected by EM visualization, whereas the periphery of U87 and other
tumor types showed a high density of collagen fibrils. These results
suggest that unassembled molecules between the fibers of the
interstitial matrix can influence the diffusion of macromolecules in
vivo just as they seem to do in vitro. In pure collagen type I gels,
these unassembled molecules can only be collagen type I, while in vivo,
these unassembled molecules may include other matrix molecules, such as
nonfibrillar collagen type I, other collagen types, or HA.
At concentrations relevant to the tumors studied, pure collagen is a major diffusive barrier and offers more hindrance than pure hyaluronan
The diffusion data attest to the ability of collagen gels at concentrations comparable to those of the tumor IM to significantly hinder diffusive transport (Fig. 2). In contrast, HA solutions at concentrations comparable to the tumors analyzed here (0.05%) pose a weaker barrier to diffusion. For 3% and 4.5% collagen gels, the diffusive barrier offered by HA (i.e., D/D0) is far less than that offered by collagen, suggesting that in tumors with these collagen concentrations (e.g., HSTS26T and U87), collagen alone can account for the diffusive hindrance in the tumor. For the lowest collagen concentration gels (1%), the barrier offered by HA is over half the barrier offered by collagen, suggesting that in tumors with this collagen concentration (e.g., LS174T) HA may have some influence on diffusive hindrance.
This finding does not apply to tissues with higher HA content,
including the tumor spheroids studied by Davies et al.
(2002) and other GAG-rich tissues, such as cartilage.
Furthermore, the pure HA solutions do not replicate possible in vivo
interactions between different species of matrix molecules (e.g.,
Turley et al., 1985), which may affect transport properties.
Collagen gels studied here pose a greater diffusive than hydraulic barrier
Data collected from several organs have indicated that
permeability is inversely correlated to collagen content
(Levick, 1987). We have found the same trend in collagen
gels. However, the permeability values in tumors did not match the data
in collagen gels quantitatively, nor did they show the qualitative
inverse correlation with collagen content. Furthermore, when the data
for collagen gels were adjusted for area fraction and tortuosity in
tumors, the permeability was higher in tumors than in gels of
comparable concentration. The differences in permeability could be due
partially to measurement techniques. Even within tumors, the confined
compression technique used by Netti et al. (2000)
predicted significantly higher hydraulic conductivity compared to the
micropipette approach (Boucher et al., 1998) and clamp
methods (Griffon-Etienne et al., 1999). The lack of
correlation between collagen and permeability observed by Netti et al.
in tumors suggests a more important contribution from other matrix molecules.
Estimates of gel permeability based on the effective medium model
matched experimental measurements of permeability only for 1% collagen
gels (Fig. 6). At greater concentrations, the diffusion-based effective
medium model values increasingly underestimated the true permeability.
In contrast, the model was reported to be accurate for agarose gels
(Pluen et al., 1999), and underestimated diffusion coefficients in various other gels, a deviation qualitatively opposite to that observed here (Phillips, 2000). In
general, discrepancies between gel measurements and effective medium
model predictions may result from model assumptions of fiber rigidity,
immobility, and homogeneity. Furthermore, the effective medium model
empirically relates two fundamentally different modes of transport
(convection and diffusion), which can be differentially regulated. The
accuracy of the effective medium prediction at low collagen
concentration and the increasing discrepancy at higher collagen
concentrations may also indicate that high concentrations of poorly
organized collagen pose a greater barrier to diffusion than to
convection. This argument is also supported by the observation that
diffusional hindrance in tumors correlates with collagen content
(Pluen et al., 2001), whereas the measured permeability
of tumors does not (Netti et al., 2000).
| |
CONCLUSIONS |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
In conclusion, our data show that collagen at physiological
concentrations presents a major barrier to molecular diffusion, especially for larger particles. Furthermore, theoretical correction of
gel diffusion data for the effects of in vivo tortuosity yielded good
agreement with in vivo measurements in tumors of comparable collagen
concentration. The diffusive hindrance data combined with imaging of
the gels and permeability measurements suggest that unassembled
collagen in the void spaces of the gel plays a role in hindering
diffusion. In vivo, this role may be played by unassembled collagen or
other matrix molecules. These findings support our hypothesis that
collagen is a major contributor to diffusive hindrance in tumors. In
addition, it suggests that in vitro gel models can be used to
investigate diffusion in tissues, with theoretical correction for
issues such as tortuosity providing the necessary bridge between the in
vivo and in vitro measurements. This work has important implications
for drug delivery in tumors and for tissue engineering, where transport
in collagen-based tissue replacements or scaffolds is an important
design consideration. Furthermore, interfering with collagen synthesis
or reducing collagen content may improve drug delivery to tumors (McKee
et al., 2001).
| |
ACKNOWLEDGMENTS |
|---|
The authors thank Dr. John M. Tarbell for the use of the permeability measurement apparatus, Dr. Robert Campbell for preparation of the liposomes used in diffusion measurements, Dr. Milind Rajadhyaksha for his advice during the development of the confocal reflectance protocol, Sylvie Roberge for her assistance in preparing samples for electron microscopy, and Mary McKee and the Massachusetts General Hospital Program in Membrane Biology for their help with the electron microscopy.
This work was supported by Outstanding Investigator Grant R35-CA56591 and Program Project Grant P01-CA-80124 from the National Cancer Institute (to R.K.J.). S.R. and E.B.B. are supported by National Institutes of Health Fellowships F32-CA83248 (to S.R.) and F32 CA88490 (to E.B.B.). T.D.M. is supported by National Institutes of Health Grant 5 T32 GM08334, Interdepartmental Biotechnology Program, Biotechnology Process Engineering Center at the Massachusetts Institute of Technology.
| |
FOOTNOTES |
|---|
Address reprint requests to Rakesh K. Jain, Department of Radiation Oncology, Massachusetts General Hospital, 100 Blossom St., Cox 7, Boston, MA 02114. Tel.: 617-726-4083; Fax: 617-724-1819; e-mail: jain{at}steele.mgh.harvard.edu.
Submitted June 25, 2001 and accepted for publication May 20, 2002.
Saroja Ramanujan's present address is Entelos, Inc., 4040 Campbell Suite #200, Menlo Park, CA 94025.
Alain Pluen's present address is School of Pharmacy and Pharmaceutical Sciences, University of Manchester, Oxford Road, Manchester, MA13 9PL, United Kingdom.
| |
REFERENCES |
|---|
|
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
CONCLUSIONS
REFERENCES
|
|---|
2 and 
1 integrins and CD44.
Cancer Res.
57:2061-2070
Biophys J, September 2002, p. 1650-1660, Vol. 83, No. 3
© 2002 by the Biophysical Society 0006-3495/02/09/1650/11 $2.00
This article has been cited by other articles:
 |
|
 |
|
  J. Henshaw, B. Mossop, and F. Yuan Relaxin treatment of solid tumors: effects on electric field-mediated gene delivery Mol. Cancer Ther., August 1, 2008; 7(8): 2566 - 2573. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. M. Ghajar, X. Chen, J. W. Harris, V. Suresh, C. C. W. Hughes, N. L. Jeon, A. J. Putnam, and S. C. George The Effect of Matrix Density on the Regulation of 3-D Capillary Morphogenesis Biophys. J., March 1, 2008; 94(5): 1930 - 1941. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Magzoub, S. Jin, and A. S. Verkman Enhanced macromolecule diffusion deep in tumors after enzymatic digestion of extracellular matrix collagen and its associated proteoglycan decorin FASEB J, January 1, 2008; 22(1): 276 - 284. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. M. Williams, W. R. Zipfel, M. L. Tinsley, and C. E. Farnum Solute Transport in Growth Plate Cartilage: In Vitro and In Vivo Biophys. J., August 1, 2007; 93(3): 1039 - 1050. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Braeckmans, K. Remaut, R. E. Vandenbroucke, B. Lucas, S. C. De Smedt, and J. Demeester Line FRAP with the Confocal Laser Scanning Microscope for Diffusion Measurements in Small Regions of 3-D Samples Biophys. J., March 15, 2007; 92(6): 2172 - 2183. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. R. Sisan, R. Arevalo, C. Graves, R. McAllister, and J. S. Urbach Spatially Resolved Fluorescence Correlation Spectroscopy Using a Spinning Disk Confocal Microscope Biophys. J., December 1, 2006; 91(11): 4241 - 4252. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. E. Fleury, K. C. Boardman, and M. A. Swartz Autologous Morphogen Gradients by Subtle Interstitial Flow and Matrix Interactions Biophys. J., July 1, 2006; 91(1): 113 - 121. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. A. Znati, M. Rosenstein, T. D. McKee, E. Brown, D. Turner, W. D. Bloomer, S. Watkins, R. K. Jain, and Y. Boucher Irradiation Reduces Interstitial Fluid Transport and Increases the Collagen Content in Tumors Clin. Cancer Res., November 15, 2003; 9(15): 5508 - 5513. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. Braeckmans, L. Peeters, N. N. Sanders, S. C. De Smedt, and J. Demeester Three-Dimensional Fluorescence Recovery after Photobleaching with the Confocal Scanning Laser Microscope Biophys. J., October 1, 2003; 85(4): 2240 - 2252. [Abstract] [Full Text] [PDF]
|
|
| |||||
) and data for LS174T, MCAIV, and U87cw (
) (a);
3% gels (
) and HSTS26T (
) (b); and 4.5% gels (
)
and U87dc (
) (c). (d) Effective tortuosity
necessary to account for discrepancy between uncorrected gel data
(Dgel) and tumor data
(DIM) as a function of tracer molecule
hydrodynamic radius. Values are calculated as 
) obtained from multiple samples and presented earlier
in Fig. 
