? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 118 Number 1 January 2008
Heparan sulfate and syndecan-1 are essential
in maintaining murine and human intestinal
epithelial barrier function
Lars Bode,1 Camilla Salvestrini,2 Pyong Woo Park,3 Jin-Ping Li,4 Jeffrey D. Esko,5
Yu Yamaguchi,1 Simon Murch,6 and Hudson H. Freeze1
1Burnham Institute for Medical Research, La Jolla, California, USA. 2Centre for Pediatric Gastroenterology, Royal Free Hospital, London, United Kingdom.
3Department of Medicine, Baylor College of Medicine, Houston, Texas, USA. 4Department of Medical Biochemistry and Microbiology, Biomedical Center,
Uppsala University, Uppsala, Sweden. 5Department of Cellular and Molecular Medicine, UCSD, La Jolla, California, USA.
6Clinical Sciences Research Institute, Warwick Medical School, Coventry, United Kingdom.
Failure to maintain intestinal epithelial barrier integrity can be
fatal. Influx of bacteria and bacterial products cause necrotizing
enterocolitis (1) and sepsis (2); excessive efflux of plasma proteins
into the intestinal lumen causes protein-losing enteropathy (PLE).
PLE develops as a symptom in ostensibly unrelated disorders,
including Crohn disease (3), multiple congenital disorders of gly-
cosylation (CDG) (4–6), and systemic lupus erythematosus (7) or
as a long-term complication of Fontan surgery to correct congeni-
tal heart malformations (8).
Emerging commonalities from clinical observations of PLE
patients alerted us to key features of PLE pathogenesis (9–11). It
is episodic, and the onset is often associated with viral infections
and increased IFN-γ levels (12, 13) as well as with a proinflam-
matory state and increased TNF-α levels (14). IFN-γ and TNF-α
disrupt tight junctions (15–17) and induce paracellular protein
leakage (9–11). PLE can result from mesenteric venous hyperten-
sion. Fontan surgery, for example, leads to general venous hyper-
tension (18). CDG-Ib patients, whose N-glycosylation is impaired
due to genetic phosphomannose isomerase deficiency, develop
hepatic fibrosis and portal hypertension (4, 5, 19). Increased pres-
sure directly causes protein leakage across epithelial monolayers
in vitro, which is further amplified upon IFN-γ/TNF-α exposure
(11). These results correlate with clinical observations in post-
Fontan patients: venous pressure increases shortly after surgery,
causing a subtle increase in intestinal protein leakage (20). How-
ever, PLE manifests only months to years after the surgery when
additional insults, usually viral infections, strike (12). Half of the
post-Fontan patients with PLE die (8), due in large part to inade-
quate therapeutic options, which are currently limited to albumin
infusions and medications that improve hemodynamics or miti-
gate inflammation, but these treatments are often accompanied
by serious side effects (21).
The most intriguing commonality in PLE patients is the specific
loss of heparan sulfate (HS) from the basolateral surface of intes-
tinal epithelial cells (IEC) during PLE episodes (6, 22, 23). Not only
HS, but also syndecan-1 (Sdc1), the predominant HS proteogly-
can (HSPG) on IEC, disappear (6). HS and Sdc1 reappear when
PLE resolves, suggesting a functional link between HS and HSPG
[HS(PG)] loss and protein leakage. HS(PG) loss in glomeruli has
been implicated in the development of proteinuria (24). Whether
HS(PG) loss also contributes to protein leakage in the intestine
is unknown. To address this question, we established an in vitro
PLE model and showed that paracellular albumin flux through
monolayers of human HT29 IEC indeed increases in the absence
of cell-associated HS (9, 11).
High-molecular-weight heparin reverses PLE in some post-Fon-
tan patients (25–27), but the basis and mechanisms are unknown.
IFN-γ and TNF-α bind to both HS and heparin (28, 29), which
may lower the local concentration of active cytokines that would
Nonstandard?abbreviations?used: AAT, α1-antitrypsin; CDG, congenital disorders
of glycosylation; dp, degree of polymerization; 2/3-DS-H, 2,3-de-O-sulfated heparin;
GAG, glycosaminoglycan; HS, heparan sulfate; HSase, heparinase III; HSPG, HS pro-
teoglycan; HS(PG), HS and HSPG; IEC, intestinal epithelial cells; PLE, protein-losing
enteropathy; rh-, recombinant human; rm-, recombinant mouse; Sdc1, syndecan-1;
TER, transepithelial electrical resistance.
Conflict?of?interest: The authors have declared that no conflict of interest exists.
Citation?for?this?article: J. Clin. Invest. 118:229–238 (2008). doi:10.1172/JCI32335.
230? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 118 Number 1 January 2008
otherwise impair intestinal epithelial integrity. In fact, we have
shown that soluble heparin compensates for loss of cell-associ-
ated HS and prevents IFN-γ/TNF-α–induced protein leakage in
vitro (9, 11). Since long-term therapy with anticoagulant heparin
has severe side effects, including bleeding, thrombocytopenia, and
osteoporosis (27, 30), an alternative therapy is needed.
Prior studies of PLE pathomechanisms and therapeutic options
were limited to in vitro cell culture models or experiments on dogs
carrying an unknown genetic defect (31, 32). Here we present what
we believe to be the first model to study intestinal protein leak-
age in mice, which allows us to control and mimic both genetic
insufficiencies and environmental insults in an in vivo setting. We
adapted well-established clinical assays (33, 34) to assess intestinal
protein leakage in mice. We found that loss of HS or Sdc1 directly
caused protein leakage and made the intestine more susceptible to
proinflammatory cytokines and increased pressure. Heparin and
non-anticoagulant 2,3-de-O-sulfated heparin (2/3-DS-H) prevent-
ed cytokine-induced protein leakage both in vitro and in mice.
Sdc1 knockdown causes protein leakage in vitro. First we assessed
whether Sdc1 loss causes protein leakage in vitro. Sdc1 siRNA
knockdown in HT29 cells reduced Sdc1 mRNA levels by 87% ± 3%
(Figure 1A) and decreased the amount of cell-associated glycos-
aminoglycans (GAGs) by 48% ± 7% (Figure 1B). In parallel, paracel-
lular protein leakage increased 1.43 ± 0.26–fold and transepithe-
lial electrical resistance (TER) decreased by 17.8% ± 6.9% (Figure 1,
B–D). Neither cell growth nor apoptosis were affected (data not
shown). Incubating HT29 cells with HS-cleaving heparinase III
(HSase) reduced cell-associated GAGs by 81% ± 3%, increased pro-
tein leakage 1.62 ± 0.19–fold, and decreased TER by 23.4% ± 6.5%.
The latter results are in accordance with our previous studies
(9, 11) and also indicate that loss of HS increases protein leakage
more than just the loss of the predominant HSPG, Sdc1.
Sdc1 knockdown amplifies TNF-α–induced protein leakage in vitro.
In the presence of HSPGs, TNF-α (20 ng/ml) increased protein
leakage 4.11 ± 0.30–fold (Figure 1E). In the absence of cell-associ-
ated HS after exposure to HSase, TNF-α increased protein leak-
age 7.00 ± 0.29–fold, showing that HS loss and TNF-α synergize.
Incubating Sdc1 knockdown cells with TNF-α increased protein
leakage 5.55 ± 0.32–fold over baseline, indicating that the effects
of Sdc1 loss and TNF-α also synergize. Again, loss of HS was more
effective than just the loss of the major HSPG Sdc1. Coincubat-
ing TNF-α with soluble heparin (25.0 μg/ml) alleviated TNF-α–
induced protein leakage.
Intestinal protein leakage is increased in Sdc1–/– or intestinal HS–defi-
cient mice. To study whether HS loss also causes intestinal pro-
tein leakage in vivo, we established 2 independent methods to
assess enteric protein loss in mice, one determining fecal loss of
51Cr-labeled albumin, the other measuring fecal levels of α1-anti-
trypsin (AAT) (10). The 2 methods gave comparable results, vary-
ing less than 10%. We applied them to mice deficient in Sdc1 or all
HS. In Sdc1–/– mice HS expression on the basolateral surface of IEC
was significantly reduced (Figure 2, A and B) and basal intestinal
protein leakage was significantly increased compared to wild-type
littermates (Figure 2C).
Since treatment of cultured mucosal cells with HSase increased
protein leakage to a greater extent than deletion of Sdc1, we next
addressed whether general loss of intestinal epithelial HS further
exacerbates protein leakage. Ext1 encodes a subunit of the copoly-
merase involved in HS formation (35). Since systemic knockout
of Ext1 is embryonic lethal (36), we inactivated the gene selec-
tively in IEC (Ext1Δ/Δ). HS expression on the basolateral surface
of IEC was further reduced compared with Sdc1–/– mice (Figure 2,
Sdc1 knockdown in HT29 cells causes protein leakage. (A–E) Sdc1-targeting siRNAs (siRNA1 and siRNA2) reduce Sdc1 mRNA level (A),
reduce cell-associated GAGs (B), increase relative albumin leakage (C), reduce TER (D), and amplify TNF-α–induced protein leakage (E), which
can be reversed with heparin. Compared with effects of HS loss after exposure to HSase or scrambled siRNA (scr. siRNA) as negative control.
White bars represent basal levels without intervention. *P < 0.05, **P < 0.01, ***P < 0.001.
? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 118 Number 1 January 2008
A and B), and basal fecal AAT levels were increased (Figure 2C).
Sdc1–/– and Ext1Δ/Δ mice did not exhibit proteinuria or hypoal-
buminemia. Gross intestinal phenotype and histology appeared
normal in both lines.
Mice (HPA-Tg) overexpressing human heparanase, which selec-
tively cleaves HS and causes a significant shortening of HS chains in
organs (37), present with proteinuria (38), and we found they also
show a 1.67 ± 0.12–fold increase in fecal AAT levels compared with
wild-type littermates (Figure 2C). Notably, protein leakage was sig-
nificantly higher in Ext1Δ/Δ and HPA-Tg mice than in Sdc1–/– mice.
TNF-α induces intestinal protein leakage in vivo, which is further enhanced
in Sdc1–/– mice. PLE onset in patients is associated with moderately
increased TNF-α levels (14), and TNF-α induces protein leakage
in vitro (9, 11). To address whether TNF-α causes intestinal pro-
tein leakage in mice, we injected wild-type mice with a single dose
of recombinant human TNF-α (rhTNF-α; 0.1 mg/kg, i.v.). These
low doses were shown not to cause inflammation or necrosis (39).
Intestinal protein leakage increased within 24 hours,
reached a maximum after 48 hours, and then slowly
decreased, reaching baseline levels after 6–7 days (Fig-
ure 3A). Recombinant mouse TNF-α (rmTNF-α) gave
similar effects as rhTNF-α (data not shown). rhTNF-α
and recombinant mouse TNF-α (rmTNF-α) have a very
similar affinity for TNFR1, but rhTNF-α does not bind
to murine TNFR2 (40), indicating that protein leakage
is mostly mediated through TNFR1 signaling.
TNF-α–induced leakage further increased in Sdc1–/–
mice. The combined effects of Sdc1 loss and TNF-α
injections were synergistic (Figure 3, A and C), consis-
tent with our in vitro results. For example, in wild-type
mice, protein leakage increased 3.25 ± 0.76–fold 48
hours after injection of rhTNF-α (Figure 3C). Theo-
retically, the individual effects of Sdc1 loss and TNF-α
presence together would predict a 3.84-fold increase
in protein leakage in Sdc1–/– mice, but we measured a
4.62 ± 0.29–fold increase, which indicated that these
factors worked synergistically.
Exogenous rhTNF-α or rmTNF-α were cleared from
mouse plasma by more than 99% within 8 hours fol-
lowing injection (data not shown; the reported half-life
is <20 min; ref. 39), and protein leakage only increased
for a few days without causing any clinical signs of
PLE, i.e., hypoalbuminemia or edema. More sus-
tained protein leakage was observed when we injected
TNF-α every 48 hours at higher concentrations (0.25 mg/kg) (Fig-
ure 3B). Effects on protein leakage progressively decreased, which
is in accordance with observations that rats develop a partial toler-
ance to daily rhTNF-α injections (39). Plasma albumin levels in the
mice remained normal, and there were no signs of edema.
IFN-γ upregulates TNFR1 and amplifies TNF-α–induced leakage,
which is further enhanced in Sdc1–/– mice. PLE onset in patients is
often associated with increased IFN-γ levels (13). We previously
showed in HT29 cells that 12 hours of exposure to IFN-γ upregu-
lates TNFR1 expression and amplifies TNF-α–induced protein
leakage (11). We now assessed the effects of IFN-γ on protein
leakage in mice. Injecting rmIFN-γ was reported to have no appar-
ent effect on mouse intestinal architecture and histology over a
wide concentration range (0.005–5.0 mg/kg) (39). We injected
rmIFN-γ (0.2 mg/kg, i.v.) and measured no increase in protein
leakage. We then assessed whether IFN-γ amplified TNF-α–
induced leakage by injecting rmIFN-γ (0.2 mg/kg, i.v.) 12 hours
Loss of Sdc1 or HS causes increased intestinal protein
leakage in mice (A) Sulfated GAG staining with colloidal
gold (left and middle columns) and HS staining with bFGF-
biotin probe (right column) in small intestinal sections from
wild-type (top row), Sdc1–/– (middle row), and Ext1Δ/Δ mice
(bottom row). The middle column is a magnification of the
left column. Original magnification, ×100 (left column); ×400
(middle column); ×200 (right column). (B) HS staining inten-
sity on basolateral surface of IEC in Sdc1–/– and Ext1Δ/Δ
mice relative to wild-type controls. (C) Intestinal protein
leakage (51Cr or AAT) in Sdc1–/–, Ext1Δ/Δ, and HPA-Tg mice
relative to respective littermate controls. All data represent
assessment in a minimum of n = 5 mice (mean ± SD).
**P < 0.01, ***P < 0.001.
232? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 118 Number 1 January 2008
prior to injecting TNF-α (0.1 mg/kg, i.v.). Intestinal protein leak-
age increased 4.17 ± 0.14–fold 48 hours after TNF-α injection,
which was significantly higher than after TNF-α exposure alone,
without prior IFN-γ injection. Calculating the additive effects of
Sdc1 loss, IFN-γ, and TNF-α, we predicted a 4.76-fold increase in
protein leakage but actually measured a 6.80 ± 0.49–fold increase
in protein leakage Sdc1–/– mice after IFN-γ and TNF-α injections,
again showing synergism. FACS analysis of SGLT1-positive
IEC revealed that IFN-γ exposure increased TNFR1 expression
2.4 ± 0.2–fold and 3.0 ± 0.4–fold in wild-type and Sdc1–/– mice
(P < 0.01), respectively (Figure 3D), showing the importance of
HS(PG) for regulating IFN-γ–induced TNFR1 expression.
Pressure induces protein leakage ex vivo and synergizes with the effects
of Sdc1 loss and cytokines. Mesenteric hypertension is common in
PLE patients (4–6, 18). Our in vitro data showed that increased
hydrostatic pressure alone causes protein leakage, which is further
amplified by HS loss, IFN-γ, and TNF-α (11). To address whether
increased pressure causes protein leakage in mice, we mounted
stripped mucosal explants (consisting of epithelial cells, lamina
propria, and muscularis mucosa but not circular and longitudi-
nal muscle layers and serosa) in Ussing chambers and applied a
hydrostatic pressure difference to the serosal side. Pressure alone
increased albumin flux 6.9 ± 1.0–fold and 10.9 ± 1.4–fold in
wild-type and Sdc1–/– mice, respectively, indicating that Sdc1 loss
increases the susceptibility to pressure-induced leakage (Figure 4).
Injecting mice with TNF-α (0.1 mg/kg, i.v.) 24 hours prior to pre-
paring explants increased flux 1.8 ± 0.4–fold and 3.8 ± 0.5–fold
in wild-type and Sdc1–/– mice, respectively, confirming that Sdc1
loss amplifies TNF-α–induced leakage. IFN-γ alone had no effect
on albumin flux but significantly increased TNF-α–induced leak-
age when administered 12 hours prior to TNF-α injection, which
was further enhanced in Sdc1–/– mice, confirming previous results.
TNF-α alone or in combination with IFN-γ exacerbated pressure-
induced protein leakage. The combination of IFN-γ, TNF-α, and
pressure increased albumin flux 15.2 ± 1.3–fold in wild-type mice
but 25.3 ± 1.8–fold in Sdc1–/– mice, again demonstrating the cen-
Sdc1–/– mice are more susceptible to cytokine-induced intestinal protein leakage. (A and B) Intestinal protein leakage (51Cr) in Sdc1+/+ and
Sdc1–/– mice in response to single (A) or multiple (B) i.v. injections of TNF-α (arrows) at 0.1 or 0.25 mg/kg. Line without symbols in A represents
predicted leakage in Sdc1–/– mice if effects of Sdc1 loss and TNF-α exposure were additive. (C) Intestinal protein leakage (AAT or 51Cr) in Sdc1+/+
and Sdc1–/– mice 48 h after exposure to TNF-α (i.v. 0.1 mg/kg), IFN-γ (i.v. 0.2 mg/kg), or a combination of both, relative to basal leakage in
Sdc1+/+ mice. Dashed lines represent predicted leakage in Sdc1–/– if effects of Sdc1 loss, TNF-α, and/or IFN-γ were additive. (D) FACS analysis
(median fluorescent activity ± SD) of TNFR1 expression in SGLT1-positive IEC from Sdc1+/+ or Sdc1–/– mice in response to IFN-γ exposure
relative to basal expression in Sdc1+/+ (which was set at 1.0; data not shown) mice. All data represent assessment in a minimum of n = 3 mice.
**P < 0.01, ***P < 0.001.
? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 118 Number 1 January 2008
tral role of Sdc1 in PLE pathogenesis. A similar fold increase in
enteric protein loss was measured in PLE patients. Mice systemi-
cally haploinsufficient for either Ext1 (Ext1+/–) or Ext2 (Ext2+/–)
also were more susceptible to IFN-γ/TNF-α– and/or pressure-
induced protein leakage, but the effects were not as prominent as
in Sdc1–/– mice.
This ex vivo system showed that weak HS(PG) expression and
IFN-γ/TNF-α, both alone and in combination, make the intesti-
nal epithelial barrier more susceptible to pressure-induced leak-
age. To study the long-term effects of increased pressure, mesen-
teric hypertension would need to be increased in vivo. Mice with
induced portal vein stenosis showed a significant increase in fecal
AAT approximately 10 days after surgery, and AAT remained ele-
vated for at least 2 weeks (data not shown).
HS(PG) loss alone does not induce paracellular leakage. HS(PG) loss
caused protein leakage and amplified IFN-γ/TNF-α–induced
protein leakage in vitro and in mice. However, how HS(PG) loss
contributes to protein leakage is unknown. To assess whether
HS(PG) loss increases paracellular leakage, we harvested mouse
intestines in lanthanum buffer followed by phosphate precipita-
tion (41). Lanthanum did not pass the junctional complex, and
the extracellular space between adjacent epithelial cells was void of
lanthanum precipitates in wild-type mice (Figure 5A) but also in
Sdc1–/– mice (Figure 5B). These results indicate that HS(PG) defi-
ciency alone has no effect on paracellular leakage. Sections from
mice exposed to IFN-γ/TNF-α showed lanthanum passing the
junctional complex and appearing in the interepithelial space and
all the way down to the basal surface (Figure 5, C–E). These results
confirm that IFN-γ/TNF-α alter junctional complex integrity and
impair intestinal epithelial barrier function.
Heparin and 2/3-DS-H alleviate cytokine-induced protein leakage. Hep-
arin injections (100–500 U/kg) mitigate PLE in some post-Fontan
patients (25–27) but have side effects mostly resulting from anti-
coagulant activity (27, 30). We screened a library of non-antico-
agulant heparin-like compounds and other GAG derivatives for
their ability to reduce IFN-γ/TNF-α–induced leakage in vitro (Fig-
ure 6A). Inhibition depended on molecular size (Figure 6A, lanes
3–7). Shorter fragments (degree of polymerization 2 [dp 2], lane 4)
had almost no effect, whereas longer fragments (dp 20, lane 7)
reduced IFN-γ/TNF-α–induced leakage by more than 50%. Low-
molecular-weight heparin (lane 3) was much less effective than
high-molecular-weight heparin (lane 2), supporting clinical data
that low-molecular-weight heparin does not mitigate PLE (26).
Heparin lost some of its activity after removal of 2-, 3-, and/or,
6-O-sulfates (lanes 8–11), reduction of the carboxyl group (lane
12), or substitution of N-sulfates with N-acetyl group (lanes 13
and 14). Other GAGs such as chondroitin, dermatan, or acharan
sulfate (lanes 15–20) also slightly reduced IFN-γ/TNF-α–induced
leakage. Sulfated cyclodextran (lane 21) or sucrose octasulfate
(lane 22) had no effect. Next to unfractionated, high-molecular-
weight heparin, non-anticoagulant 2/3-DS-H (lane 9) was most
efficient in reducing IFN-γ/TNF-α–induced leakage.
We then assessed whether heparin or 2/3-DS-H also reduce
IFN-γ/TNF-α–induced protein leakage in mice by giving daily
i.v. injections of either heparin or 2/3-DS-H. Three days after the
initial injection, we administered either TNF-α alone or a combi-
nation of IFN-γ and TNF-α, as described above. Daily injections
of heparin or 2/3-DS-H continued for 7 days. Low doses of either
heparin (100 U/kg, 0.7 mg/kg) or 2/3-DS-H (0.7 mg/kg) reduced
cytokine-induced intestinal protein leakage by more than 50%
(Figure 6, B and E). Heparin doses 5-fold higher (500 U/kg, 3.5
mg/kg) completely prevented leakage (Figure 6, C and F) in wild-
type mice. In Sdc1–/– mice, higher 2/3-DS-H doses (3.5 mg/kg)
completely prevented TNF-α–induced leakage (Figure 6D) and
reduced IFN-γ/TNF-α–induced leakage by more than 85% (Figure
6G). Administration of either heparin or 2/3-DS-H alone for 10
consecutive days had no adverse effects on mice and did not alter
basal intestinal protein loss.
Our in vitro and in vivo results show that HSPGs are essential for
maintaining the intestinal epithelial barrier function. Progressive
loss of HSPGs increases basal protein leakage. The greater the HS
loss, the higher the leakage. This deficiency exacerbates cytokine-
and pressure-induced intestinal protein leakage, but heparin and
non-anticoagulant 2/3-DS-H prevent protein leakage, providing a
rationale for heparin therapy in PLE patients.
Although HSPG loss appears to play a central role in PLE
pathogenesis, the etiology of HSPG loss in PLE patients remains
unknown. We hypothesize that one or more environmental insults
Sdc1 loss and Ext1 or Ext2 haploinsuffi-
ciency increase protein leakage ex vivo.
Albumin leakage (mean ± SD) through
stripped mouse mucosal explants mount-
ed in Ussing chambers. Significances cal-
culated compared with wild-type mice with
the same interventions. All data represent
assessment in a minimum of n = 6 mice.
*P < 0.05, **P < 0.01, ***P < 0.001. n.d.,
234? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 118 Number 1 January 2008
such as infections, inflammation, or increased mesenteric pres-
sure collide with genetic insufficiencies in pathways associated
with HSPG biosynthesis, trafficking, or degradation (10). Here
we tested this hypothesis by using an extreme approach of geneti-
cally ablating either Sdc1 or intestinal epithelial Ext1, which is
unlikely to occur in PLE patients. However, even mice haploinsuf-
ficient for either Ext1 or Ext2 also show an increased susceptibil-
ity for stress-induced protein leakage. Thus the risk of developing
PLE could increase if an individual had several haploinsufficient
mutations affecting overall HSPG expression. Environmental
insults may then further exacerbate HSPG loss. TNF-α, for exam-
ple, downregulates HSPG expression (42) and also induces HSPG
shedding (43). Inducing systemic hypertension in rats reduces
HSPG expression in the glomeruli and causes proteinuria (44). We
also have evidence that increased pressure reduces Sdc1 mRNA
levels in HT29 cells (our unpublished results). Therefore, low
levels of HSPG expression in genetically susceptible individuals
may be further diminished below a critical threshold by cytokines
and pressure. In return, reduced HSPG expression amplifies the
effects of cytokines and pressure, creating a vicious cycle of more
dramatic HSPG loss, breakdown of the intestinal epithelial bar-
rier, and, eventually, PLE.
However, why HSPG loss alone increases intestinal epithelial
leakage remains elusive. Here we show by electron microscopic
examination on lanthanum-bathed intestinal tissues that HSPG
loss alone does not increase paracellular flux between adjacent
epithelial cells. If this is the case, how does protein leak across the
intestinal epithelium if the junctional complex integrity remains
intact? Watson et al. asked a slightly different question (45): How
does the epithelial barrier remain intact despite continuous cell
shedding? They showed that an impermeable substance com-
pletely fills the void left by shedded cells and speculated that this
sealing substance could be HS secreted by neighboring epithelial
cells and/or myofibroblasts (45). HS(PG) deficiency would result
in an inability to seal the gaps and, as a consequence, cause leak-
age. Further studies are needed to test this hypothesis and show
whether HS is indeed the sealing agent.
From a therapeutic perspective, the vicious cycle involving HSPG
loss, cytokines and pressure can be interrupted at multiple steps by
addition of heparin or 2/3-DS-H, compensating for the loss of cell-
associated HS and restoring the IFN-γ/TNF-α–quenching effect.
Soluble heparin binds to both IFN-γ and TNF-α, just as cell-associ-
ated HS does (28, 29), and blocks IFN-γ/TNF-α–induced protein
leakage in vitro (9, 11) and, as we show here, in mice. This may be
one explanation for the curious beneficial effects of heparin injec-
tions in some post-Fontan patients with PLE (25–27). However,
heparin has undesirable side effects due to its anticoagulant activ-
ity (27, 30). Partially removing heparin’s 2- and 3-O sulfates reduc-
es its anticoagulant activity by about 90%, while other biological
activities remain intact (46). Here we show that 2/3-DS-H, similar
to unmodified heparin, prevents IFN-γ/TNF-α–induced protein
leakage in vitro and in mice. Its greatly reduced anticoagulant
activity may mean that it can be used safely at much higher doses
than unmodified heparin. Clinical trials will be needed to assess
whether 2/3-DS-H can be considered a safe and effective therapeu-
tic option for patients suffering and dying from PLE.
Measurement of protein flux in vitro. Human intestinal epithelial HT29 cells
(ATCC no. HTB-38) were grown in Dulbecco’s modified Eagle’s medium
(Irvine Scientific) supplemented with 10% fetal calf serum (HyClone), peni-
cillin, streptomycin, and l-glutamine. Cells were grown on semipermeable
inserts (1.0 μm pore size; Becton Dickinson) for 5 days until they reached
confluence (9). Integrity of the monolayer before and after interventions
was monitored by measuring TER with an epithelial voltohmmeter and
STX2 electrodes (World Precision Instruments). TER in the untreated
confluent HT29 monolayer was defined as 100% after correcting for the
resistance of the semipermeable membrane in tissue culture media but
without cells. To assess protein leakage across the monolayer, albumin-
FITC (400 μg/ml; Sigma-Aldrich) was added to the inserts. Albumin-FITC
concentration in the well (across the monolayer) was measured after 1 h
using a spectrophotometer. Albumin flux through the untreated confluent
monolayer was defined as 1.0.
Sdc1 siRNA knockdown and induction of HS loss. HT29 cells were transfected
with Sdc1 siRNA1 (CCAUUCUGACUCGGUUUCUTT), Sdc1 siRNA2
(GCCAAGGUUUUAUAAGGCUTT), or scrambled siRNA (all Ambion) as
negative control using siLentFect Lipid (BioRad). Sdc1 mRNA levels were
measured by quantitative PCR 5 days after transfection. To induce loss
of cell-associated HS, HT29 cells were incubated with HSase (0.6 mU/ml;
Sdc1 loss alone does not increase paracellular leakage. Electron
micrographs of mouse intestinal epithelium bathed in lanthanum. Lat-
eral intercellular spaces were void of lanthanum phosphate precipi-
tates (arrows) in wild-type (A) and Sdc1–/– mice (C) without cytokine
exposure but filled with lanthanum phosphate precipitates in wild-type
(B), Sdc1–/– (D), and Ext1Δ/Δ mice (E) after IFN-γ/TNF-α exposure.
Right column is a magnification of boxed areas in the left column.
Scale bars: 5 μm.
? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 118 Number 1 January 2008
Heparin and 2/3-DS-H alleviate protein leakage in vitro and in mice. (A) Albumin leakage (mean ± SD) through HT29 monolayers relative to
untreated cells. Maximum leakage (white bar) was induced by incubating cells with heparinase (HS loss), IFN-γ (10 ng/ml, 24 h), and TNF-α
(2 ng/ml, 12 h). Cytokines were coincubated with heparin-like compounds or other GAG derivatives at 2.5 μg/ml or 25.0 μg/ml. Heparin (lane 2)
and 2/3-DS-H (lane 9) were most effective in alleviating cytokine-induced protein leakage (arrows). Lane 1, HS; lane 2, high-molecular-weight
heparin (unfractionated); lane 3, low-molecular-weight heparin; lane 4, sized heparin, dp 2; lane 5, sized heparin, dp 8; lane 6, sized heparin, dp
14; lane 7, sized heparin, dp 20; lane 8, 2,6-de-O-sulfated heparin; lane 9, 2/3-DS-H; lane 10, 6-O-desulfated heparin (chemical desulfation);
lane 11, 6-O-desulfated heparin (enzymatic desulfation with endosulfatase [HSulf2]); lane 12, carboxyl-reduced heparin; lane 13, fully N-acety-
lated heparin; lane 14, fully O-sulfated N-acetylated heparin; lane 15, chondroitin sulfate; lane 16, fully O-sulfated chondroitin sulfate; lane 17,
dermatan sulfate; lane 18, fully O-sulfated dermatan sulfate; lane 19, fully O-sulfated hyaluronic acid; lane 20, archaran sulfate; lane 21, sulfated
cyclodextran; lane 22, sucrose octasulfate. (B–G) Intestinal protein leakage in Sdc1+/+ (B, C, E, and F) and Sdc1–/– mice (D and G) assessed by
in vivo 51Cr labeling. Mice were injected daily with low (100 U/kg, 0.7 mg/kg) (B and E) or high doses (500 U/kg, 3.5 mg/kg) of heparin (C and F),
2/3-DS-H (C, D, F, and G), or PBS as control. Three days following the first injections (t = 0), intestinal protein leakage was induced by injection
of either TNF-α (i.v., 0.1 mg/kg) (B–D) or IFN-γ (i.v., 0.2 mg/kg) and TNF-α (i.v., 0.1 mg/kg, 12 h after IFN-γ) (E–G). PBS was used as a control.
All data represent assessment in a minimum of n = 4 mice (mean ± SD).
236?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 118 Number 1 January 2008
Sigma-Aldrich) for at least 1.5 h. To suppress reappearance of cell-associ-
ated HS, cells were incubated with β-xyloside (p-nitrophenyl-β-d-xylopy-
ranoside, 100 μM; Sigma-Aldrich), which competes out GAG chain syn-
thesis on core proteins.
Measurement of cell-associated GAGs. HT29 cells were incubated with
Na235SO4 (100 μCi/ml) for 8 h. Cells were harvested in 4 M guanidine
HCl/50 mM acetic acid/1% Triton X-100 and diluted with 50 volumes of
6 M urea/50 mM acetic acid/1% Triton X-100. Radiolabeled proteoglycans/
GAGs were purified by anion exchange chromatography on DE53 columns
(Whatman) as described previously (9). Radioactivity was determined by
Mice. Sdc1–/– mice were provided by Merton Bernfield (Harvard Medical
School, Boston, Massachusetts, USA) (47) and bred for more than 12 gen-
erations onto a C57BL/6 genetic background. Sdc1–/– mice were genotyped
as previously described (47). Homozygous transgenic mice overexpressing
human heparanase (HPA-Tg) were generated and genotyped as described
(38). The HPA-Tg mice were on a mixed genetic background. Results
from these mice were compared with those of control animals with the
same genetic background. Ext1+/– and Ext2+/– mice (48), backcrossed on a
C57BL/6 genetic background, were provided by Jeffrey Esko (University of
California, San Diego, La Jolla, California, USA) and genotyped as previ-
ously described (48). Mice with an intestine-specific Ext1 knockout were
generated by crossing Ext1flox/flox mice provided by Yu Yamaguchi (Burnham
Institute for Medical Research, La Jolla, California, USA) (49) with Villin-
Cre mice (Vil-Cre; The Jackson Laboratory) expressing Cre recombinase
under the control of the 12.4-kb Villin promoter, which drives exclusively
in the intestinal epithelium starting at day E12.5. (50). Penetrance of Cre
expression in these tissues is almost 100% (50). Both Ext1flox/flox and Vil-
lin-Cre mice were on a C57BL/6 genetic background. Ext1Δ/Δ mice were
genotyped as previously described (49). To exclude age and gender effects,
all experiments were performed on 3- to 4-month-old male mice. Unless
otherwise indicated, wild-type littermates were used as controls. All mouse
work was approved by the Animal Research Committee at the Burnham
Institute for Medical Research.
Measurement of protein flux through stripped mouse mucosal explants. Mice
were euthanized, and small intestines were immediately excised and rinsed
in oxygenized Ringer solution containing 10 mM glucose. Intestines were
opened along the mesenteric lining. Serosa, longitudinal, and circular
muscle layers were stripped off. The remaining mucosal layer (including
muscularis mucosa, lamina propria, and epithelial cells) was mounted into
Ussing chambers. Both sides of the tissue were bathed in Ringer’s solu-
tion gassed with 95% O2 and 5% CO2. Hydrostatic pressure difference was
induced by adding additional buffer volume to the compartment facing
the serosal tissue side (10). Protein leakage across the tissue was deter-
mined by measuring flux of albumin-FITC from the serosal to the mucosal
side. Albumin flux through tissues from wild-type mice without further
intervention was defined as 1.0. Prior to preparing mucosal explants, some
mice were injected with either rmIFN-γ (0.2 mg/kg, i.v., RND), rhTNF-α
(0.1 mg/kg, i.v., RND), or a combination of both.
Intestinal protein leakage in mice. Two independent methods were used to
assess intestinal protein leakage in mice. (a) 51CrCl3 (300,000 cpm) was
injected in the tail vein to label plasma albumin. Radioactivity of 24-h
feces samples was measured with a gamma counter (10). Fecal excretion
correlates with intestinal protein leakage (34). (b) AAT levels in 24-h feces
samples were determined by competitive ELISA (10).
Immunohistochemistry: sulfated GAG staining. Sulfated GAG distribution
was localized on formalin-fixed tissues as previously reported (22, 51).
Briefly, cationic colloidal gold (polylysine gold; Biocell International) was
suspended 1:100 in PBS at pH 1.2 and applied to the tissue sections for 1 h
at room temperature. Bound polylysine gold was visualized with a silver
enhancer. Tissue sections were counterstained with Mayer’s hemalum and
mounted with Aquamount.
Immunohistochemistry: HS staining with biotinylated FGF. FGF was biotinyl-
ated as previously described (52). Briefly, FGF2 (0.5 mg, E. coli recombi-
nant material; Selective Genetics Inc.) was protected with heparin (0.4
mg) in 0.2 M HEPES buffer (pH 8.4) and mixed with biotin hydrazide
(long arm, water-soluble, 40 μg; Pierce Biotechnology) in a final vol-
ume of 0.2 ml. After 2 h at room temperature, 40 μl of 10 mg/ml gly-
cine solution was added to stop the reaction. The sample was diluted
with 30 ml of 20 mM HEPES buffer (pH 7.4) containing 0.5 M NaCl and
0.2% BSA and loaded onto a 1-ml column of heparin-Sepharose CL-6B
(Amersham Pharmacia Biotech). The column was washed with 30 ml of
buffer and eluted with 2.5 ml of solution adjusted to 3 M NaCl. The
sample was desalted on a PD-10 column (Amersham Pharmacia Biotech)
equilibrated with 20 mM HEPES buffer (pH 7.4) containing 0.2% BSA.
Inclusion of 125I-FGF2 (~4 Å, ~105 cpm; ref. 48) showed that the over-
all recovery of material was 30%–40%. Paraffin-embedded small intes-
tine tissue sections were labeled with the FGF-biotin probe (40 μg/ml,
overnight, 4°C), as previously described (53). Bound factor was detected
with HRP-conjugated streptavidin (1:500, 30 min at room temperature;
Jackson ImmunoResearch Laboratories). To determine staining density,
ImageJ software was used on digital photomicrographs, using plug-ins
for color deconvolution (A.C. Ruifrok, Netherlands Forensic Institute,
The Hague, The Netherlands) and region-of-interest densitometry
(J. Kuhn, University of Texas, Austin, Texas, USA; Anthony Padua, Duke
University, Greensboro, North Carolina, USA). Mean density scores were
obtained for each slide from multiple villous tip epithelial basolateral
membranes. A staining density index was derived from reciprocals of
light transmission, minus background constant, but all statistical analy-
sis was performed on unprocessed data.
Electron microscopy. Mouse small intestines (5–10 mm) were dis-
sected and fixed for 3 h at room temperature in a buffer containing 4%
paraformaldehyde, 1.5% glutaraldehyde (in 0.1 M cacodylate buffer), and
1% lanthanum nitrate hexahydrate, followed by an additional 5 h in a
buffer containing 3% glutaraldehyde (in 0.1 M cacodylate buffer) and 1%
lanthanum nitrate hexahydrate. To precipitate lanthanum, tissues were
incubated overnight with a 0.2-M phosphate buffer at 4°C. Tissues were
postfixed for 2 h at room temperature in 1% OsO4 with 0.15% ferricyanide
in 0.1 M cacodylate buffer, washed for 1 h in 0.1 M cacodylate buffer,
and dehydrated in 50%, 70%, 90%, and 100% ethanol for 15 min each. Tis-
sues were cleared in 2 changes of propylene oxide for 15 min and then
impregnated overnight in Epon/Araldite resin. Afterward, samples were
embedded and polymerized at 60°C. Thick sections (0.5–2.0 μm) were
stained in toluidine blue for general evaluation. Afterward, thin sections
(60 nm) were cut and mounted on parlodion-coated copper slot grids.
Sections were stained with uranyl acetate and lead citrate and analyzed at
FACS analysis and TNFR1 expression. Mouse IEC were harvested as previ-
ously described (54). Cells were stained with anti-TNFR1–specific (Santa
Cruz Biotechnology Inc.) and IEC-specific anti-SGLT1 antibodies (Alpha
Diagnostic) and analyzed by flow cytometry (Becton Dickinson). SGLT1-
positive cells were gated, and median fluorescence activity for TNFR1 was
recorded. Data are expressed as median fluorescence activity ± SD relative
to cells from wild-type mice without intervention.
Statistics. Data are expressed as mean ± SD unless otherwise stated.
Groups were compared by the Student’s 2-tailed paired or unpaired t test.
A value of P < 0.05 was considered statistically significant. Unless otherwise
stated, in vitro results represent a minimum of 3 evaluations, in vivo results
are obtained from a minimum of n = 3 mice, and ex vivo data is obtained
from a minimum of n = 6 mice.
?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 118 Number 1 January 2008
We thank Robert J. Linhardt (Rensselaer Polytechnic Institute,
Troy, New York, USA) for providing us with a heparin-like com-
pound library, Steven Rosen and Kenji Uchimura (UCSF, San
Francisco, California, USA) for providing endosulfatase-treated
heparin, and Lianchun Wang (University of Georgia, Athens,
Georgia, USA) for providing us with 2/3-DS-H. We thank Kim E.
Barrett and Declan McCole (UCSD) for their support with pre-
paring stripped mucosal explants and with the Ussing chamber
experiments. We gratefully acknowledge the late Merton Bernfield
(Harvard Medical School, Boston, Massachusetts, USA) for pro-
viding us with the Sdc1–/– mouse line. We thank Israel Vlodavsky
(Bruce Rappaport Faculty of Medicine, Technion, Israel) for the
HPA-Tg mice. We thank Malcolm Woods (Director Core Micros-
copy Facility, The Scripps Research Institute, La Jolla, California,
USA) for his excellent support with electron microscopy. We also
thank Wilfred Charbono and Adriana Charbono for their excellent
technical assistance with mice, Edward Monosov and Guillermina
Garcia for technical assistance with immunohistochemistry, and
Mien Nguyen for technical assistance with mouse genotyping.
This work was supported by NIH grants (R01 DK065091 and R21
HL 078997, to H.H. Freeze; R01 HL57345 to J.D. Esko) and grants
from Children’s Hearts Fund (to H.H. Freeze) and Deutsche Forsc-
hungsgemeinschaft (BO-2488/1-1, to L. Bode).
Received for publication April 6, 2007, and accepted in revised
form October 17, 2007.
Address correspondence to: Hudson H. Freeze, Burnham Institute
for Medical Research, 10901 North Torrey Pines Road, La Jolla,
California 92037, USA. Phone: (858) 646-3142; Fax: (858) 713-
6281; E-mail: email@example.com.
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