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Regular Articles |
From the Cardiovascular Research Institute and Department of
Anatomy,*
University of California, San Francisco,
California; and the Edwin L. Steele
Laboratory,
Department of Radiation Oncology,
Massachusetts General Hospital and Harvard Medical School,
Boston, Massachusetts
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Of particular functional importance, tumor vessels are unusually leaky.15-20 Vessel leakiness correlates with histological grade and malignant potential of tumors.21 Furthermore, the leakiness of tumor vessels can result in extravasation of plasma proteins and even erythrocytes and may facilitate the traffic of tumor cells into the bloodstream and the formation of metastases.19,22-24 Understanding the basis for this abnormal permeability is key to the effective delivery of chemotherapeutic agents to tumor cells and to monitoring the efficacy of antiangiogenic therapy.
Evidence for increased endothelial permeability of tumor vessels has come from many sources. Extravasation of soluble tracers such as radioisotopes, albumin, fibrinogen, dextran, horseradish peroxidase, Lissamine green, and ferritin has been demonstrated in experimental tumors.16,25-28 Studies of the extravasation of blood-borne particulates such as colloidal carbon and sterically stabilized (stealth) liposomes has revealed functional pore sizes as large as 2 µm in tumor vessels.29-31 Scanning electron microscopy (EM) has shown the extravasation of intravenously (i.v.) injected plastic casting media in some tumors.3,32 The leakiness of tumor vessels has also been documented by magnetic resonance imaging of i.v. injected contrast media.33,34 Intratumoral hemorrhage is a manifestation of vessel leakiness that can range from scattered extravasated erythrocytes to blood lakes (also called hemorrhagic lakes or vascular lakes), which are large collections of erythrocytes surrounded by tumor cells.22,35-38 Alternatively, blood lakes could be abnormal vascular channels connected to the bloodstream.39,40
Vessel leakiness in tumors has been attributed to highly active angiogenesis and microvascular remodeling, but the structural basis and mechanism of the leakage are unclear. Intercellular gaps, transendothelial holes, vesiculo-vacuolar organelles, and endothelial fenestrae are all reportedly present in the endothelium of tumor vessels.19,41 However, the relative frequency and contribution of these to vessel leakiness have not been studied extensively. By comparison, the mechanism of vessel leakiness in inflammation is much better characterized. Most studies point to the involvement of transiently open intercellular gaps in leakage from inflamed vessels.42,43 Under some conditions transcellular holes may also play a role.44
The purpose of the present study was to identify structural abnormalities in the endothelium of tumor blood vessels that could explain their leakiness. In particular, we sought to determine whether the vessels have intercellular gaps or transcellular holes similar to those found in leaky vessels of inflamed tissues. We also determined whether blood lakes in tumors are collections of erythrocytes that have extravasated from leaky blood vessels or abnormal vascular channels connected to the bloodstream.
We reasoned that structural abnormalities should be most easily detected in tumors with very leaky vessels. Therefore, our strategy was to focus on a tumor, MCa-IV mouse mammary carcinoma, that is known to have blood vessels on the high end of the range of leakiness for tumors, as evidenced by a functional pore size of 1200 to 2000 nm.31 Less leaky Shionogi mammary tumors (pore size 200–380 nm), LS174T colon carcinomas (pore size 400–600 nm), and normal mouse mammary glands were used for comparison. In addition, insulinomas in RIP-Tag2 transgenic mice, which have a bloody appearance, were used to examine the nature of blood lakes in tumors.45,46 After tissues were fixed by vascular perfusion, we used CD31 immunohistochemistry, binding of intravascular lectins, uptake of fluorescent cationic liposomes, and vascular labeling with the particulate tracer Monastral blue to confirm the identity of blood vessels and to minimize the chance of misinterpreting abnormal vessel-like spaces in tumors. This approach also made it possible to distinguish between intravascular erythrocytes and extravascular erythrocytes. Then the luminal surface of vessels exposed in Vibratome sections was examined by scanning EM to identify intercellular gaps, transcellular holes, or other potential sites of leakage across the endothelium. The scanning EM observations were confirmed by transmission EM.
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Two-millimeter cubes of MCa-IV mouse mammary carcinomas were implanted under the dorsal skin of male C3H mice (25–30 g) in the animal facility of University of California San Francisco or the Edwin L. Steele Laboratory, Department of Radiation Oncology, Massachusetts General Hospital, as previously described.31 The tumors grew to a diameter of 8 to 12 mm over 14 to 20 days.31 For comparison, Shionogi tumors (mouse testosterone-dependent male mammary carcinomas) and LS174T human colon adenocarcinomas were grown subcutaneously to a diameter of about 8 mm over 14 to 20 days in male SCID mice.31 Spontaneous pancreatic islet cell tumors were studied in RIP-Tag2 transgenic mice at 8 to 14 weeks of age.47 Blood vessels were also examined in normal mammary glands of C3H mice. Mice were anesthetized with ketamine (87 mg/kg) plus xylazine (13 mg/kg) injected together intramuscularly. The chest was opened rapidly, and the vasculature was perfused for 2 minutes at a pressure of 120–140 mmHg with fixative from a cannula (blunt 18 gauge needle) inserted into the ascending aorta via an incision in the left ventricle. The cannula was immediately clamped in place (time from heart incision to perfusion <15 seconds). The composition of the fixative varied with the purpose of the experiment (see below). The fixative was not preceded by a buffer or saline rinse. The right atrium was incised to create a route for the fixative to exit. With this approach, the vasculature was washed free of blood and preserved in an open state; the only blood remaining in tumors was extravasated erythrocytes. All experimental procedures were approved by the Committees on Animal Research of University of California San Francisco and the Massachusetts General Hospital.
Scanning Electron Microscopy
For scanning EM, tissues were fixed by vascular perfusion of 0.5% glutaraldehyde and 1% paraformaldehyde in 0.075 mol/L sodium cacodylate buffer, pH 7.4. Tumors were removed, immersed in 2.5% glutaraldehyde in cacodylate buffer for a minimum of 2 hours, and embedded in 10% agarose. The luminal surface of blood vessels in tumors was exposed by cutting Vibratome sections 100 µm in thickness. The sections were rinsed with cacodylate buffer, immersed in cacodylate buffered 2% tannic acid for 24 hours, rinsed, and then immersed in 2% OsO4 in 0.1 mol/L cacodylate buffer for 2 hours at 4°C.48 After dehydration with ethanol, sections were infiltrated with 100% t-butanol and freeze-dried under vacuum. Sections were sputter-coated with a 4- to 16-nm-thick layer of gold-palladium alloy. The luminal surface of vessels was photographed with a JEOL JSM-840A or Hitachi S-2380N scanning electron microscope.
Brightfield and Fluorescence Microscopy
For routine histology, tissues were fixed by vascular perfusion of 0.5% glutaraldehyde and 1% paraformaldehyde in phosphate-buffered saline (PBS), embedded in glycol methacrylate resin (JB-4, Polysciences, Warrington, PA), sectioned at 2 µm thickness, and stained with hematoxylin and eosin. Specimens were examined with a Zeiss Axiophot microscope equipped for brightfield, differential interference contrast, and epifluorescence.
Tumor vasculature was made visible by labeling with biotin- or fluorescein isothiocyanate (FITC)-labeled Lycopersicon esculentum lectin (100 µg in 100 µl 0.9% NaCl, Vector Laboratories, Burlingame, CA) injected into the femoral vein of anesthetized mice, and 2 minutes later fixative containing 0.5% glutaraldehyde and 1% paraformaldehyde in PBS was perfused through the vasculature. Tumors were embedded in 10% agarose and cut with a Vibratome or cryostat (100 µm thickness). Biotin-labeled lectin was visualized by incubation overnight with avidin-biotin peroxidase complex (ABC; Vector Laboratories) diluted 1:250 in phosphate-buffered saline containing 0.3% Triton-X 100 (PBS/Triton) followed by diaminobenzidine (DAB; Sigma Chemical Company, St. Louis, MO), dehydration, and mounting in Permount (Fisher Scientific, San Francisco, CA).49 The detergent permeabilized tissues, increasing the penetration of reagents into the 100-µm sections. FITC-labeled lectin in sections mounted in Vectashield (Vector) was visualized by fluorescence microscopy.
Tumor blood vessels were also visualized by their uptake of fluorescent cationic liposomes, which have a high affinity for endothelial cells in tumors and other sites of angiogenesis.12,50 Cationic liposomes approximately 50 nm in diameter, consisting of 55% 1,2-dioleoyl-3-trimethylammonium-propane (DOTAP), 44% cholesterol, and 1% rhodamine-DHPE (1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine) were prepared by sonication and suspended in 5% dextrose.50 Lipids were purchased from Avanti Polar Lipids (Alabaster, AL). The liposome suspension (1440 nmol total lipid in 150 µl of 5% dextrose) was injected into a femoral vein 20 minutes before vascular perfusion of fixative (0.5% glutaraldehyde and 1% paraformaldehyde in PBS). Some mice received i.v. injections of both liposomes and FITC-labeled lectin to assess colocalization of the two markers. Vibratome or cryostat sections were cut, mounted, and viewed by fluorescence microscopy.
The question of whether blood lakes in tumors of RIP-Tag2 mice were connected to the vasculature was addressed by labeling blood vessels with fluorescent lectin or cationic liposomes injected i.v. as described above. Alternatively, the particulate tracer Monastral blue (1 µl/g body weight of 3% suspension of copper phthalocyanine blue pigment BW-431-P, Chemicals and Pigments Department, E. I. duPont deNemours and Company, Wilmington, DE) was injected into a femoral vein of anesthetized mice, and 10 or 20 minutes later fixative (0.5% glutaraldehyde and 1% paraformaldehyde in PBS) was perfused through the vasculature. The pancreas and its tumors were removed as a block, and sites of Monastral blue labeling were localized by brightfield microscopy in whole mounts mounted in Permount.
Endothelial cells were localized by CD31 (PECAM) immunohistochemistry in tumors fixed by vascular perfusion of 4% paraformaldehyde in PBS and embedded in 10% agarose (Sigma) or infiltrated overnight with 30% sucrose and frozen. Sections 100 µm in thickness, cut with a Vibratome or cryostat, were incubated at room temperature for 12 to 15 hours in rat anti-mouse CD31 monoclonal antibody (Pharmingen, San Diego, CA) diluted 1:500 followed by 6 hours in biotin- or Cy3-labeled donkey secondary anti-rat IgG antibody (Jackson ImmunoResearch, West Grove, PA) diluted 1:200 in PBS/Triton. Sections were then processed and viewed as for lectin staining.
Transmission EM
Tumors were fixed by vascular perfusion of aldehyde fixative as for scanning EM. Specimens measuring approximately 1 x 3 mm were cut from 100-µm Vibratome sections, treated with OsO4 and uranyl acetate, dehydrated, and embedded in epoxy resin.51 Sections 0.5 µm in thickness were stained with toluidine blue for light microscopy, and sections 50–100 nm in thickness were stained with lead citrate and examined with a Zeiss EM-10 electron microscope.
Morphometry and Statistical Analysis
Diameters of 300 blood vessels in six MCa-IV tumors were measured
in video images of methacrylate sections with a digitizing tablet
interfaced with a computer.51
Approximately 700 cells
lining blood vessels of MCa-IV tumors in 5 mice were photographed by
scanning EM at both low and high magnifications as needed for different
measurements. The proportions of branched and unbranched lining cells
were determined in 20 vessels having a total visible surface area of
220,000 µm2
(magnification, 500 to 1000x). The
luminal surface area and perimeter of a sample of 25 unbranched cells
and 25 branched cells in 35 vessels were measured using a digitizing
tablet (magnification, 2000 to 5000x). The shape index of cells was
calculated from their area A and perimeter P by
the equation 4
x
A/P2, where the shape index
of a circle is 1. The shape index decreases as the profile becomes more
elongated or irregular. The area and perimeter of 100 intercellular
openings between the 700 lining cells (magnification, 2000 to 10,000x)
were measured, as were the 8 transcellular holes identified along with
the openings. Intercellular openings that exposed the plasma membrane
of an underlying cell were excluded because they were assumed not to be
pathways for extravasation. Area measurements were used to calculate
the total amount of luminal surface examined (291,000
µm2) and the combined areas of the openings
(292 µm2) and holes (2.3
µm2). The diameter of intercellular openings
and transcellular holes was estimated from their areas (2 *
SQRT(A/)) or perimeters
(P/2), with the assumption that they were roughly
circular. Values are presented as means ± SE. The significance of
differences between means was assessed by Student’s t-test.
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Several different methods of identification were used to verify
that the blood vessel-like structures in MCa-IV tumors were not
artifactual spaces, cysts, collections of extravasated erythrocytes, or
other abnormalities. By all methods, the blood vessels were found to be
emptied of blood and preserved in an open state by the vascular
perfusion of fixative. In hematoxylin-and-eosin-stained sections of
MCa-IV tumors, blood vessels were abundant, thin-walled, and surrounded
by 100-µm thick sleeves of tumor cells interspersed by regions of
necrosis (Figure 1A)
. The vessels were
variable in size (diameter 39 ± 7 µm, range 8–220 µm, 300
vessels in 6 tumors). Larger vessels were conspicuous and easily
identified by their size and their thin but distinct wall (Figure 1B)
,
but small vessels predominated, with 48% being 25 µm or less in
diameter (Figure 2)
. Most large vessels
were separated from tumor cells by a perivascular space as wide as 20
µm containing stromal cells and extracellular matrix, yet some were
much closer to tumor cells (Figure 1B)
. Collections of extravasated
erythrocytes were rare in MCa-IV tumors but were common in islet cell
tumors in RIP-Tag2 mice (see below). MCa-IV tumors were surrounded by a
capsule. Most of these features, including the blood vessels,
perivascular sleeves of tumor cells, regions of necrosis, and capsule,
were also visible by scanning EM (Figure 1C)
.
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. ( The term "vessel
lining cells" is used instead of "endothelial cells" to describe
the cells in contact with blood in tumor vessels because of the
possibility that some of these cells may not be endothelial cells.)
Green lectin and red liposomes were colocalized in the abundant small
vessels in the capsule at the tumor surface as well as on vessels of
all sizes within the tumor (Figure 1, D and E)
. (We do not categorize
tumor vessels as arterioles, capillaries, or venules because the
vessels lack the structural characteristics of normal vasculature that
make these terms meaningful.) The labeled vessels had an irregular
shape and widely variable diameter and branching pattern (Figure 1, G and H)
, which was quite unlike the uniform, smooth, cylindrical contour
of the normal microvasculature. Extravasation of the lectin blurred the
labeling of some vessels (Figure 1, D and G)
and resulted in green
fluorescence of scattered regions of extravascular cells. No evidence
of liposome extravasation was detected. In addition to the red coating
of the lining cells, some liposome fluorescence was punctate,
consistent with an endosomal location.
The pattern of CD31 immunoreactivity in MCa-IV tumors (Figure 1F)
was
more complex and extensive than the vessel pattern shown by lectin
staining or cationic liposome binding (Figure 1, D and E)
. Although the
overall arrangement of vessels was similar, more structures had CD31
immunoreactivity than were stained by the other methods (Figure 1I)
. In
particular, thin CD31-immunoreactive sprout-like filaments, without a
lumen detectable by lectin staining (Figure 3A)
, radiated from some vessel walls into
the perivascular sleeves of tumor (Figure 3B)
. Similar sprouts were
evident by scanning EM, and again no lumen was visible (Figure 3C)
.
Many of these traversed the 100-µm thickness of the sleeves of tumor
tissue, and some contacted other vessels (Figure 3, D and E)
. CD31
immunoreactivity colocalized with lectin staining on the vessel lining
but not on the sprouts that radiated from vessel walls into the tumor
(Figure 3, G and H)
, indicating that these sprouts did not have a lumen
connected to the blood circulation. CD31-immunoreactive sprouts were
particularly conspicuous when visualized by DAB staining (Figure 1, F and I)
.
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Blood vessels of normal mammary glands examined by scanning EM
were lined by flat endothelial cells that were closely apposed
to one another, had a smooth contour, and were demarcated by slightly
raised margins (Figure 4A)
. The
endothelial cells formed a continuous, uniform monolayer, as has been
described in the microcirculation of other organs.48,51
The longest cytoplasmic projections of endothelial cells measured less
than 1 µm.
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-D).
The vessel lining cells in MCa-IV tumors probably represented a
continuous spectrum of shapes but tended to fall into two groups,
designated unbranched cells and branched cells, based on whether they
had long cytoplasmic projections. An average of 86 ± 2% of the
vessel surface was covered by unbranched cells (20 vessels in 5 MCa-IV
tumors). These cells differed from endothelial cells in vessels of
normal mammary glands because of their variable size and irregular
shape (Figure 4, A and B)
. Some of the cells had a protruding nucleus,
ruffled margins, prominent cytoskeleton, or short stubby processes
(Figure 4B)
.
The remaining 14 ± 2% of the luminal surface of vessels in
MCa-IV tumors was covered by branched cells (Figure 4, C and D)
. These
cells typically were multipolar, and some had projections as long as 50
µm (Figure 4E)
. The cells were conspicuous because of their highly
irregular shape, abnormal thickness, partial detachment, and long
cytoplasmic projections that overlapped other lining cells (Figure 4E)
or bridged the vessel lumen (Figure 4F)
. Cells corresponding to the
branched cells were also evident by light microscopy (Figure 5A)
. Some branched cells that were
stained with lectin and CD31 immunoreactivity extended well into the
vessel lumen (Figure 5, B and C)
.
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Defective Endothelial Monolayer
Branched cells did not form an intact monolayer, as judged by
scanning EM. Instead, the cells formed two, three, or perhaps more
incomplete layers in which the cells next to the lumen covered
underlying cells, and cell projections extended between the layers
(Figure 4, E and F)
. Projections of some branched cells spanned the
luminal surface of one or more neighboring cells (Figure 5D)
or bridged
the vessel lumen (Figure 5E)
. Other cell projections penetrated between
or even through adjacent cells (Figure 5F)
. Regions of branched cells
between the projections formed tunnels of various size, length, and
complexity where they arched over other lining cells (Figures 4E, 4F, and 5D)
. The arches created intercellular openings in the vessel wall
(see below).
Defects in the endothelial monolayer and lining cells with bizarre
branched shapes were also found by scanning EM in vessels of Shionogi
testosterone-dependent male mouse mammary carcinomas, LS174T human
colon adenocarcinomas, and spontaneous pancreatic islet cell tumors in
RIP-Tag2 mice. Some lining cells formed arches, transluminal processes,
or multiple layers (Figure 5, G
-I). However, the abnormalities were
less severe than those in MCa-IV tumors.
Tumors in RIP-Tag2 mice not only had vessels with a defective lining
but also contained extravasated erythrocytes that gave them a bloody
appearance even after vascular perfusion of fixative (Figure 6A)
. Some of the extravasated
erythrocytes were located in discrete blood lakes (Figure 6A)
. Unlike
blood vessels, blood lakes were not labeled by fluorescent lectin or
cationic liposomes injected into the vasculature (Figure 6B)
.
Similarly, Monastral blue injected i.v. was not found within the lumen
or wall of blood lakes but did label some vessels at the tumor surface.
Thus, there was no evidence that the blood lakes were directly
connected to the bloodstream. Although the lakes resembled large,
sack-like blood vessels or sinusoids by light microscopy (Figure 6C)
,
it was obvious by transmission EM that they were lined by tumor cells,
not endothelial cells (Figure 6D)
. Indeed, some erythrocytes were
immediately adjacent to tumor cells (Figure 6E)
. The extravascular
location of erythrocytes in RIP-Tag2 tumors was unambiguous near blood
vessels where the vessel lumen and lining cells were visible (Figure 6, F and G)
. Scanning EM examination showed in a complementary way that
the vasculature in these tumors was lined by cells that resembled
endothelial cells (Figure 7A)
and that
blood lakes were bordered by tumor cells (Figure 7, B and C)
.
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. Lining
cells in contact with the vessel lumen were heterogeneous in appearance
(Figure 7, D and E)
and had many features not found in normal
endothelial cells. Some regions of lining cells were tightly apposed
and joined by junctions (Figure 7E)
, whereas others were separated from
one another (Figure 7F)
. The cells varied in thickness from ~0.1 µm
in attenuated regions to as much as 4 µm near the nucleus (Figure 7E)
, and some had irregular finger-like projections (Figure 7G)
. The
lining cells had a normal looking nucleus and cytoplasm rich in rough
endoplasmic reticulum and mitochondria, typical of metabolically active
rather than apoptotic cells (Figure 7, E and F)
.
Vessel lining cells had a basement membrane that blended into the
extracellular matrix around the stromal cells located between vessels
and tumor cells (Figure 7, F and G)
. Tumor cell clusters were also
bordered by basement membrane. When multiple layers of flattened cells
were located between the vessel lumen and tumor cells, vessel lining
cells were not readily distinguished from pericytes or stromal cells
(Figure 7G)
. The basement membrane-like extracellular matrix, which
consisted in some regions of multiple layers and in others of a solid
matrix (Figure 7, D, F, and G)
, contributed to the difficulty in
distinguishing endothelial cells, pericytes, stromal cells, and tumor
cells by transmission EM.
Intercellular Openings
Round or oval openings were present between lining cells of many
vessels in MCa-IV tumors examined by scanning EM (Figure 8, A
-C). Diameters calculated from the
areas of 100 openings in 5 tumors averaged 1.7 ± 0.2 µm
(mean ± SE; range, 0.3–4.7 µm). These values were conservative
estimates because they tended to minimize the pore size. The size was
skewed slightly toward smaller openings, with a median of 1.5 µm
(Figure 9)
. Diameter estimates from
perimeters (2.2 ± 0.2 µm; range, 0.3–5.4 µm) gave the
maximum diameter openings could achieve, regardless of their shape.
Intercellular openings were visible between 29% of the cells, assuming
100 openings among the 700 cells examined and the involvement of two
cells per opening. These openings constituted approximately 0.1% of
the luminal surface examined (292 µm2
of
291,000 µm2). Basement membrane was visible
through five of the openings (Figure 8A)
; the remainder appeared dark
by scanning EM, and the underlying structures could not be identified.
Narrow slit-like spaces, similar to the oblique slits that form between
endothelial cells of inflamed vessels,48
were found
between many cells, but the size and patency of these pathways were
difficult to judge; therefore, they were excluded from the
measurements. Openings in the lining layer were also evident by
transmission EM (Figure 8D)
.
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Eight transcellular holes were found along with the 100
intercellular openings in the sample of 700 vessel lining cells in
MCa-IV tumors examined by scanning EM (Figure 8, B and C)
. The diameter
calculated from the area of transcellular holes was 0.6 ± 0.1
µm (range, 0.2–0.9 µm; Figure 9
). The interior was dark and
unidentifiable in seven of the holes, but in one, the plasma membrane
of an underlying cell was visible, so it is unclear whether it was a
route for extravasation. The eight holes occupied a total area of 2.3
µm2, which was 0.8% of the area represented by
the 100 intercellular openings present in the same sample and 0.0008%
of the luminal surface area.
Fenestrae with diaphragms approximately 50 to 80 nm in diameter were
visible by transmission EM in thin regions of cells lining MCa-IV tumor
vessels (Figure 8E)
. Clusters of fenestrae were visible by scanning EM
(Figure 8F)
in 30% of a sample of 55 lining cells; 59% of these
fenestrated cells were branched cells.
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Pathways for Plasma Leakage in Tumors
The main goal of this study was to identify the anatomical pathway for extravasation of particles as large as 2 µm from tumor vessels.30,31 Our scanning and transmission EM observations revealed multiple defects in the lining of MCa-IV tumor vessels and identified three pathways that have been implicated in plasma leakage: openings between the vessel lining cells, holes through the lining cells, and endothelial fenestrae. Of these, intercellular openings are the most likely to explain the unusual leakiness of the MCa-IV tumor vessels. The 1.7 µm average diameter of the intercellular openings corresponds well to the 1.2 to 2.0 µm functional pore cutoff size measured by the extravasation of sterically stabilized (stealth) liposomes in these tumors.31 The openings were more than 10 times as numerous as transcellular holes, and their combined surface area was over 100 times that of transcellular holes. By comparison, transcellular holes in vessels of MCa-IV tumors had an average diameter of only 0.6 µm.
Data from the present study confirmed the presence of endothelial fenestrae in vessels of MCa-IV tumors.31 Fenestrae are present in many other tumors as well as in vessels exposed to vascular endothelial growth factor and in some types of normal vessels.5,52,53 However, endothelial fenestrae are unlikely candidates for the extravasation of 2-µm tracer particles because of their much smaller diameter (~50–80 nm) and diaphragm with channels of < 6 nm diameter.54
The morphological data obtained not only match the functional pore size but also fit well with previous evidence of intercellular openings in the vessels of MCa-IV tumors31 and are consistent with many other reports of open junctions or endothelial gaps, spaces, or discontinuities in tumor vessels.31,41,52,53,55-57 Further support for the involvement of intercellular openings was the use of multiple complementary methods without particulate tracers that are reported to alter permeability.58 This evidence does not, however, exclude the participation of pinocytotic vesicles or vesiculo-vacuolar organelles in transvascular transport across tumor vessels.25,59,60 Indeed, it has been suggested that vesiculo-vacuolar organelles give rise to transendothelial holes.44,61 The process of macropinocytosis, used by dendritic cells to internalize soluble antigens constitutively,62 is another possibility, but we are not aware of evidence that this process can translocate 2-µm particles across the endothelial barrier.
Hemorrhage is another sign of tumor vessel leakiness.22,35,36,38,63 Many extravasated erythrocytes were present individually and in blood lakes in RIP-Tag2 tumors, but few were present in MCa-IV tumors. Blood lakes were a conspicuous feature of enlarged "angiogenic" islets and tumors in the pancreas of RIP-Tag2 mice and were readily seen with the naked eye, although these red spots have previously been interpreted as angiogenic blood vessels that form at a particular stage of tumorigenesis.45,46 We found that the blood lakes were lined by tumor cells and did not communicate directly with the vasculature. Our finding that the lakes did not contain cationic liposomes or Monastral blue, which have short half-lives in the bloodstream, suggests that blood lakes form when vessel wall integrity is transiently disrupted. This periodic hemorrhage may be linked to sprouting angiogenesis driven by growth factors that predispose to bleeding.13,22,64
The presence of extravasated erythrocytes in tumors has further significance. A recent publication argues that blood vessels of malignant eye tumors known as uveal melanomas are formed by tumor cells instead of endothelial cells.40 The argument is based in part on ultrastructural evidence of erythrocytes located next to tumor cells, which were presumed to line vascular channels. However, a more likely explanation, suggested by other studies of uveal melanomas39,65 and the present findings, is that most erythrocytes in contact with tumor cells are not moving through the tumor but rather have left the circulation and are stagnant, at least over the time scale of 30 minutes or so. Studies using long half-life tracers or intravital microscopy may help to determine the time course of extravasation and fate of these cells.
Intercellular Openings versus Intercellular Gaps in Tumor Vessels
We used the term intercellular openings to distinguish the large, irregular spaces between branched lining cells of tumor vessels from intercellular gaps that form at endothelial cell borders in inflamed vessels.43 The intercellular openings of tumor vessels were much larger than the uniform ~0.5-µm intercellular gaps produced by inflammatory mediators.48,51 Also, the intercellular openings in tumor vessels had no obvious structural features that could result in their closure, unlike rapidly reversible endothelial gaps in inflamed vessels, which are spanned by fingerlike projections that may assist in their closure. Exactly how intercellular openings form in tumor vessels is not understood, although growth factors like vascular endothelial growth factor and angiopoietin-2 may contribute.13,64 Also, endothelial cells in tumor vessels have a greater motility and more rapid turnover (higher mitotic and apoptotic rates) than normal endothelial cells.66-71 As a result, intercellular junctions and extracellular matrix attachments may not form normally, leading to impaired monolayer formation and barrier function.
Identity of Lining Cells in Tumor Vessels
In the present study, we have cautiously referred to the lining cells of tumor vessels, rather than assuming that these cells are endothelial cells. Blood vessels in MCa-IV tumors were lined by a distinct layer of CD31-immunoreactive cells and some of the cells had fenestrae. Therefore, these cells appeared to be endothelial cells, albeit abnormal. Although no unstained regions were detected, we recognize that it would be difficult to identify scattered CD31-negative cells in our tissue preparations. Also, despite the apparent homogeneity of CD31 staining, the lining cells were obviously morphologically heterogeneous, with unbranched and branched phenotypes, and the presence of other cell types such as tumor cells, pericytes, or stromal cells cannot be excluded.
In some tumors, blood vessels have been reported to have a discontinuous or absent endothelial cell lining, so that blood flows directly past exposed tumor cells.1,40,52,72 It has also been claimed that tumor cells can be incorporated as structural elements of the vessel wall and contribute to a cellular mosaic of lining cells.1,73 Even though our results are consistent with abnormal endothelial cells constituting the lining of MCa-IV tumors, our immunohistochemical studies did not provide conclusive information about the other tumors. Definitive identification of the lining cells, particularly the branched phenotype, awaits unambiguous ultrastructural localization with the appropriate cell markers.
There is no consensus on the single best marker of endothelial cells. Immunohistochemical markers, such as von Willebrand factor and P-selectin, and properties such as uptake of acetylated low-density lipoprotein and lectin staining are not uniformly distributed in endothelial cells of the normal microvasculature,50,74 and their characteristics in abnormal vessels are incompletely understood. For this reason, we labeled tumor blood vessels by several different methods, and analyzed the similarities and differences in the results. CD31-immunoreactive cells in MCa-IV tumors appeared to be restricted to vessel lining cells and sprouts emanating from them, consistent with what has been observed in other tumors.75 However, some tumor cells have been reported to have CD31 immunoreactivity.40,76,77
Like CD31 immunoreactivity, i.v. injected L. esculentum lectin identified vessel lining cells in tumors, but the lectin extravasated from some tumor vessels and stained scattered cells in the stroma. Cationic liposomes also labeled the lining cells of tumor vessels and produced a sharper image of tumor vessels than lectin staining because of the limited extravasation.12 Cationic liposomes intensely labeled all tumor vessels, consistent with our previous observations that it is not just the newly formed vessels in tumors that avidly take up these liposomes. Pre-existing parts of the tumor vasculature behave the same as vessels undergoing remodeling and angiogenesis.12 This finding is consistent with our scanning EM observations suggesting that the lining cells of tumor vessels are much more dynamic than normal endothelial cells.
Transluminal Processes, Sprouts, and Other Abnormalities of Vessels
Cells with cytoplasmic projections that bridged the vessel lumen are reminiscent of endothelial cells in vessels undergoing intussusceptive growth in which transluminal bridges eventually partition the vessel lumen.78,79 Our finding that the number of structures with CD31 immunoreactivity exceeded the number of vessels labeled by lectin or cationic liposomes probably reflects the presence of endothelial cell sprouts without lumens that are growing into the tumor75,80 or regions of intermittent perfusion.81 These sprouts may be vessel precursors.
Scanning EM provided detailed, informative views of the luminal surface of vessels, but gave little information about the deeper layers of the vascular wall. Observations from lectin staining, CD31 immunohistochemistry, uptake of fluorescent cationic liposomes, and transmission EM confirmed the multilayered nature of the vessel lining and the heterogeneity of the lining cells. Vessels in the tumors examined were not enveloped by layers of normal smooth muscle cells. Further studies will be needed to determine the number and distribution of pericytes in vessel walls and the role of pericytes in permeability defects in tumor vessels.
Clinical Implications of Tumor Vessel Leakiness
The presence of a defective endothelial monolayer with large anatomical pathways for extravasation has important implications for viral and nonviral gene therapy and other macromolecular treatments of cancer that depend on the leakiness of tumor vessels for their efficacy.82 Also, studies using dynamic magnetic resonance imaging suggest that tumor vessel permeability is correlated with malignancy in an experimental tumor model.21 The observation that endothelial cells of some vascular sprouts are not in contact with the vessel lumen indicates that blood borne antiangiogenic agents may not have direct access to all endothelial cells in tumors.
Vascular endothelial growth factor and the angiopoietins are critical regulators of the balance between vascular growth, maturation, and regression, and the leakiness associated with the defective endothelial monolayer of tumor vessels.13 If vessel leakiness is a functional abnormality that helps tumors survive, therapeutic reversal of the leak through agents targeted to endothelial cells may be clinically beneficial.83
In conclusion, we have shown that blood vessels in tumors known to be leaky have a defective endothelial monolayer. Although some tumor vessels have relatively normal lining cells, others are lined by branched, overlapping, disorganized, and loosely interconnected cells. Most of the lining cells appear to be abnormal endothelial cells, but the presence of other cell types cannot be excluded. Spaces between the poorly connected, branched lining cells are the most likely candidates for the pathway for extravasation of particles as large as 2 µm from tumor vessels.
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Supported in part by National Institutes of Health grants HL-24136 and HL-59157 from the National Heart, Lung and Blood Institute (to D. M.) and R35-CA-56591 from the National Cancer Institute (to R. J.) and by a grant from MBT Munich Biotechnology GmbH in Munich, Germany (to D. M.). D. M., J. M., and G. T. have equity positions in Munich Biotechnology.
H. H.’s present address: Department of Anatomy, Niigata University School of Medicine, Asahimachi-dori 1, Niigata 951-8510, Japan.
Accepted for publication January 6, 2000.
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