intestinal epithelial tight junctions as targets for enteric bacteria-derived toxins
TRANSCRIPT
www.elsevier.com/locate/addr
Advanced Drug Delivery Reviews 56 (2004) 795–807
Intestinal epithelial tight junctions as targets for enteric
bacteria-derived toxins
Alessio Fasanoa,b,c,*, James P. Natarob,c
aDivision of Pediatric Gastroenterology and Nutrition, School of Medicine, University of Maryland, Baltimore, MD, USAbMucosal Biology Research Center, School of Medicine, University of Maryland, Baltimore, MD 21201, USA
cCenter for Vaccine Development, School of Medicine, University of Maryland, Baltimore, MD, USA
Received 6 October 2003; accepted 3 November 2003
Abstract
The application of a multidisciplinary approach to study bacterial pathogenesis, along with the recent sequencing of entire
microbial genomes have made possible discoveries that are changing the way scientists view the bacterium–host interaction.
Today, research on the molecular basis of the pathogenesis of infectious diarrheal diseases of necessity transcends established
boundaries between microbiology, cell biology, intestinal pathophysiology, and immunology. Novel multidisciplinary
approaches led to the discovery of new bacteria–host cell interactions involving signals regulating intestinal permeability
through the modulation of cell cytoskeleton and intercellular tight junctions (TJ). A century ago, TJ were conceptualized as a
secreted extracellular cement forming an absolute and unregulated barrier within the paracellular space. Biological studies of the
past several decades have shown that TJ are dynamic structures subjected to structural changes that dictate their functional
status under a variety of developmental, physiological, and pathological circumstances. To meet the many diverse physiological
challenges to which the intestinal epithelial barrier is subjected, TJ must be capable of rapid and coordinated responses. This
requires the presence of a complex regulatory system that orchestrates the state of assembly of the TJ multiprotein network.
Many pathogenic bacteria exploit this system to accomplish their pathogenic strategies by ultimately modulating intestinal
permeability.
D 2004 Elsevier B.V. All rights reserved.
Keywords: Enterotoxins; Bacteria; Tight junctions; Paracellular pathway; Intestinal permeability
Contents
1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7962. Tight junctions, a key barrier . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7963. Structure of tight junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 796
0169-409X/$ - see front matter D 2004 Elsevier B.V. All rights reserved.
doi:10.1016/j.addr.2003.10.045
* Corresponding author. Mucosal Biology Research Center, School of Medicine, University of Maryland, 685 W. Baltimore St., HSF
Building, Room 465, Baltimore, MD 21201, USA. Tel.: +1-410-328-0812; fax: +1-410-328-1072.
E-mail address: [email protected] (A. Fasano).
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807796
3.1. Membrane proteins associated with tight junctions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7973.2. Peripheral membrane proteins associated with tight junctions . . . . . . . . . . . . . . . . . . . . . . . . 7983.3. The cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7983.4. Protein–protein interactions between transmembrane proteins,
cytoplasmic plaque proteins and the actin cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . 7984. Toxins affecting the enterocyte TJ/cytoskeleton complex . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 799
4.1. Toxins affecting the cell cytoskeleton . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7994.1.1. Clostridium difficile toxins A and B . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7994.1.2. Clostridium botulinum toxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7994.1.3. Clostridium botulinum C3 toxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8004.1.4. Escherichia coli cytotoxic necrotizing factors 1 and 2 (CNF) . . . . . . . . . . . . . . . . . . . 800
4.2. Toxins that destroy TJ structural elements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8004.2.1. Bacteroides fragilis toxin (BFT), a zinc-dependent metalloprotease toxin . . . . . . . . . . . . . 8004.2.2. Serine protease autotransporters of enterobacteriaceae . . . . . . . . . . . . . . . . . . . . . . . 8014.2.3. Vibrio cholerae hemagglutinin protease. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 802
4.3. Toxins that affect TJ competency by stimulating host signaling events . . . . . . . . . . . . . . . . . . . 8024.3.1. Zonula occludens toxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8024.3.2. Clostridium perfringens enterotoxin (CPE) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8044.3.3. Vibrio cholerae RTX (‘repeats-in-toxin’) toxin . . . . . . . . . . . . . . . . . . . . . . . . . . . 804
5. Concluding remarks and future directions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 804
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 804
1. Introduction
Microorganisms represent the first species of
living organisms that populated our planet and will
probably continue to survive well beyond the extinc-
tion of the human race. Their distinguishing charac-
teristics (small size, concise deployment of genetic
information, and ability to survive in highly varied
circumstances) contribute to their manifest virtuosity
in adapting to a changing environment. To be a
successful enteric, non-invasive pathogen, a micro-
organism has to be a good colonizer, compete
effectively for nutrients, and to be able to interact
with the target eukaryotic cell in order to induce
secretion of water and electrolytes. Since the basic
metabolism of enteric pathogens and commensals is
largely the same, it follows that pathogens must
possess highly specialized attributes, which enable
them to activate one or more eukaryotic intracellular
pathways leading to intestinal secretion. This cross
talk between enteric pathogens and the host intestine
may be affected by either invasion or elaboration of
toxins. This article is focused on the growing num-
ber of discovered enterotoxins that have been de-
scribed to exert their pathogenic effect by targeting
the cell cytoskeleton/tight junctions (TJ) complex.
2. Tight junctions, a key barrier
A key function of the intercellular junction complex
between neighboring intestinal epithelial cells (enter-
ocytes) is the formation of selective barriers that permit
the generation andmaintenance of tissue compartments
with distinct compositions. Individual enterocytes are
joined to each other by a specialized complex consist-
ing of TJ (zonula occludens, ZO), adherens junctions,
gap junctions, and desmosomes [1]. TJ represent the
major barrier within the paracellular pathway [2].
Evidence now exists that TJ, once regarded as static
structures, are in fact dynamic and readily adapt to a
variety of circumstances. The adaptive mechanisms
and specific regulation of TJ are areas of active inves-
tigation, and they still remain incompletely understood.
3. Structure of tight junctions
The actual structure of the TJ has been studied
extensively (Fig. 1). Freeze fracture electron micros-
copy reveals that these contacts, which encircle the
apical side of the lateral surface of each cell, are
continuous strand-like transmembrane structures
which interact with similar structures of adjacent cells.
Fig. 1. Model for components of the TJ. The transmembrane proteins occludin and the claudins are anatomically and functionally connected
with the cell cytoskeleton via the junctional complex. This complex comprises a series of proteins, including ZO-1, ZO-2, and p130 (ZO-3).
Other proteins, such as cingulin, 7H6, rab13, rho, and ras, are located further from the cell membrane. However, they seem also involved in the
regulation of TJ permeability.
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807 797
The interactions define the paracellular permeability
characteristics. A number of proteins are associated
with TJ.
3.1. Membrane proteins associated with tight
junctions
The first protein found to be associated with TJ,
occludin, was identified in 1993 [3]. Occludin is
composed of four transmembrane domains, two
extracellular domains, and a long cytoplasmic car-
boxyl-terminal tail. Electrophoresis evidence suggest
that the phosphorylated form of occludin appears to
be the major form located within the TJ, whereas the
less phosphorylated forms are found in the baso-
lateral membrane and in the cytosol. Knockout
experiments using murine embryo stem cells suggest
that occludin is not the only component responsible
for TJ competency [4]. Another family of TJ mem-
brane proteins, the claudins, was recently identified
[5]. Of the different claudins that have been de-
scribed, claudins 2–5 are mainly expressed in the
intestine [6]. The functional importance of claudins
in forming fibrils was demonstrated by using a
claudin-11 knockout mice model in which a com-
plete loss of TJ fibrils in Sertoli cells was observed.
It is believed that each claudin possesses selective
permeability properties. Similar to occludin, some of
the claudins of neighboring cells interact to form
intercellular connections, while other members of the
claudins do not [7]. It is also very interesting that
some claudins have additional functions. Claudin-3
and Claudin-4 were both identified to function as
receptors for Clostridium perfringens enterotoxin
(CPE) [8,9]. A third membrane protein, junction-
associated membrane protein (JAM) was identified
by raising monoclonal antibodies against endothelial
cells [10]. JAM has only one putative transmem-
A. Fasano, J.P. Nataro / Advanced Drug D798
brane domain, and the extracellular portion of JAM
contains two domains with intra-chain disulfide bonds,
which is typical of immunoglobulin-like loops of the V-
type. JAM mediates homotypic cell–cell adhesion
[10]. It remains unknown if JAM could form a func-
tional barrier to prevent the free flux of ions and small
solutes.
3.2. Peripheral membrane proteins associated with
tight junctions
ZO-1, ZO-2, and ZO-3 are cytoplasmic proteins
associated with TJ. ZO’s form a complex on the
cytoplasmic side of TJ. ZO-1 is a f 220 kDa
peripheral membrane protein that is localized in the
immediate vicinity of the plasma membrane of TJ in
both epithelial and endothelial cells [11]. ZO-1 has
been demonstrated to interact with the actin cyto-
skeleton through fodrin [12]. Other peripheral pro-
teins, called ZO-2 and ZO-3 with molecular masses
of 160 and 100 kDa, respectively, have been iden-
tified as ZO-1 binding proteins [13,14]. Sequence
analysis shows that ZO-1, ZO-2 and ZO-3 are
members of the large family of membrane-associated
guanylate kinase (MAGUK) proteins. MAGUK pro-
teins share several structural motifs, including vari-
able numbers of PDZ domains, one src homology 3
(SH3) region, and one guanylate kinase (GUK)
homology region. The PDZ domains of these pro-
teins appear to interact with the C-terminal cytoplas-
mic tail of transmembrane proteins, and this is
believed to be the mechanism of ZO-1–occludin
interaction. Several other peripheral membrane pro-
teins have also been localized to the TJ, including
cingulin, 7H6, rab 13, Gai� 2, and protein kinase C
(PKC) [15]. Another protein named symplekin has
been described that is not only associated with TJ,
but can also be localized to the nucleus [16]. In cells
that do not form TJ, symplekin appears to be
localized only in the nucleus. ZO-1 also can be
localized to the nucleus, but unlike symplekin, ZO-
1 can be found in the nucleus under growing con-
ditions and not in differentiated epithelial cells [17].
This pattern of dual localization for these TJ com-
ponents suggests that beside regulation of paracellu-
lar permeability, these structures might also be
involved in the regulation of gene expression, cell
growth, and differentiation [15].
3.3. The cytoskeleton
There is now a large body of evidence that struc-
tural and functional linkage exists between the actin
cytoskeleton and the TJ complex of absorptive cells
[18–20]. The actin cytoskeleton is composed of a
complicated meshwork of microfilaments whose pre-
cise geometry is regulated by a large cadre of actin-
binding proteins. The architecture of the actin cyto-
skeleton appears to be critical for TJ function. Most of
the actin is positioned under the apical junctional
complex where myosin II and several actin-binding
proteins, including a-catenin, vinculin, and radixin
have been identified [21]. Myosin movement along
actin filaments is regulated by ATP and phosphoryla-
tion of the regulatory light chain by Ca2 +/calmodulin-
activated myosin light chain kinase [21]. In several
systems, increases in intracellular Ca2 + can influence
phosphorylation of myosin regulatory light chain and
increase contraction of perijunctional actin and para-
cellular permeability [22]. Increased permeability has
been also linked to PKCa-dependent polymerization
of actin filaments strategically located to dictate TJ
competency [23].
3.4. Protein–protein interactions between transmem-
brane proteins, cytoplasmic plaque proteins, and the
actin cytoskeleton
The transmembrane proteins, occludin and claudin,
interact with each other and with other proteins of the
TJ complex. Occludin binds to itself, the ZO proteins,
and actin [24]. Likewise, claudins can associate with
themselves, occludin, and in some circumstances,
with other claudins [24]. In addition, most of the
claudin tails, including claudin 1, end in YV, a
sequence reminiscent of a PDZ-binding motif [24].
Indeed, a direct interaction between the first PDZ
domains of ZO-1, ZO-2 and ZO-3 with a glutathione
S-transferase (GST)-fusion protein encoding the eight
COOH-terminal amino acids in the tail of claudins 1–
8 has been recently demonstrated [25]. The role these
protein–protein interactions play in the organization
of the transmembrane proteins is unclear. In the case
of occludin, the ZO-1 binding region can act as a
signal for occludin localization [22]. Localization
mediated through adhesive interactions with cytoplas-
mic proteins may also apply to the claudins [22]. The
elivery Reviews 56 (2004) 795–807
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807 799
role of these protein–protein interactions in TJ com-
petency remains to be established.
4. Toxins affecting the enterocyte TJ/cytoskeleton
complex
Given the key function of intestinal TJ in regulat-
ing trafficking of water and molecules between envi-
ronment and host, it is not surprising that some
bacterial toxins have evolved to exploit this function
as part of their pathogenic arsenal. What is remark-
able, however, is the breadth and complexity of
strategies developed by enteric bacteria to affect
intestinal permeability. The following section outlines
the better-characterized examples of enteric toxins
affecting TJ competency.
4.1. Toxins affecting the cell cytoskeleton
Four toxins produced by Clostridium spp. and a
toxin produced by certain Escherichia coli strains
have been demonstrated to affect intestinal TJ by
modifying either Rho guanosine triphosphatases
(GTPases) or actin. These toxins have been reviewed
extensively [26–30] and will only be discussed
briefly here.
4.1.1. Clostridium difficile toxins A and B
C. difficile has emerged as the most important
pathogen causing the syndrome of antibiotic-associ-
ated colitis [31]. The virulence of this pathogen is
dependent on its elaboration of two related toxins
TxA and TxB. These toxins are among the largest
monomeric toxins described, with molecular weights
of 308,000 for TxA and 270,000 for TxB. Despite the
fact that TxA has traditionally been referred to as an
enterotoxin and TxB as a cytotoxin [31], they both
exert a cytotoxic effect in vitro. Both TxA and TxB
are glucosyltransferases and use uridine diphosphate
(UDP)-glucose as a substrate to inactivate, by mono-
glucosylation, members of the Rho family of small
GTPases at Thr37, an amino acid residue located
within the putative effector domain of the Rho pro-
teins [32]. Rho GTPases regulate a variety of cyto-
skeleton-dependent cellular functions, such as cell
adhesion and motility, growth-factor-mediated signal-
ing, cellular transformation, and induction of apopto-
sis [33]. The dramatic effects of TxA and TxB on
tissues and cells, including cytoskeletal depolymer-
ization, increased intestinal permeability and diarrhea,
cellular retraction and rounding, disruption of cell
adhesion and chemotaxis, and activation of apoptosis
[34], have been traditionally related to the TxA- and
TxB-dependent inactivation of the Rho proteins.
However, more recent findings seem to suggest that
these toxins also activate Rho-independent pathways.
In animal experiments, TxA induces hemorrhagic
fluid secretion, an inflammatory response and mark-
edly damages ileal and colonic epithelium, whereas
TxB appears inactive. However, TxB is a potent
cytotoxin when tested in cultured cells. Recent experi-
ments examining the action of TxA and TxB on
human colon in vitro revealed that both toxins stim-
ulated a decrease in intestinal barrier function [35]
and, surprisingly, toxin B was more potent than toxin
A. Additional studies of polarized intestinal epithelial
cells in vitro revealed that both toxins diminished
barrier function of the cells without inducing cytotox-
icity. However, the time course of resistance changes
and the pattern of F-actin changes in the cells were
distinct, suggesting that the toxins bind to distinct
receptors, differ in their ability to stimulate intracel-
lular signaling pathways and/or traffic differently in
intestinal epithelial cells. Additional studies of the
mechanism of action of TxA confirm that the cellular
response to toxin A is more complex than previously
hypothesized [36]. TxA localizes to mitochondria
within minutes of cellular exposure and prior to the
onset of Rho glucosylation, which occurs at a later
time [37]. Activation of multiple pro-inflammatory
pathways (e.g. release of reactive oxygen species,
activation of primary sensory neurons, stimulation of
interleukin-8 (IL-8) production) occurs rapidly and the
pathophysiologic response to TxA is attenuated by
inhibitors directed at suppressing the inflammatory
response [38].
4.1.2. Clostridium botulinum toxins
The C2 toxin induces intestinal secretion second-
ary to tissue damage [29,39]. Its role in human
disease is unknown. The C2 toxin consists of a
binding subunit (105 kDa) and an active subunit
(55 kDa) that confers the biologic activity of the
toxin. The C2 toxin exhibits substrate specificity by
ADP-ribosylating G (monomeric) actin at Arg177,
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807800
leading to loss of actin ATPase activity. ADP-ribo-
sylation of G-actin by the C2 toxin prevents its
polymerization into F-actin and, over time, results
in depolymerization of cellular F-actin, possibly
leading to the loosening of intercellular TJ.
4.1.3. C. botulinum C3 toxin
The C3 toxin ADP-ribosylates RhoA, B and C (but
not Rac or Cdc42) at Asp41 within the GTPase
effector region (reviewed in Ref. [40]). As a result,
actin filaments are disassembled in a fashion similar to
that described for C. difficile toxins. When tested on
polarized intestinal epithelial (T84) cells, actin local-
ized to the apical perijunctional ring was disas-
sembled, leading to the dissociation of the TJ
protein ZO-1 from the perijunctional complex [41].
Following these changes in protein–protein interac-
tion, the TJ barrier function was perturbed, as man-
ifested by changes of transepithelial resistance (TER)
and increased passage of the paracellular marker
dextran [41].
4.1.4. E. coli cytotoxic necrotizing factors 1 and 2
(CNF)
E. coli producing CNF-1 has been associated with
human disease including enteritis, urinary tract infec-
tions and prostatitis whereas E. coli producing CNF-2
has been associated with enteric disease of farm
animals [42–44]. The amino acid content of the two
toxins indicates that they are closely related, with 85%
identical and 99% conserved residues. Both toxins are
cell-associated and it is hypothesized that they are
delivered to host cells by the bacteria, possibly via a
Type III secretion system. The CNF toxins deamidate
Gln63 (into glutamic acid) of Rho or Gln61 of Rac and
Cdc42 (also members of the Rho GTPase protein
family). This modification blocks GTPase activity
(i.e. preventing the hydrolysis of GTP to GDP) lock-
ing the Rho proteins in their GTP-bound activated
state, and resulting in permanent activation of the
proteins [43]. In intestinal epithelial cell monolayers,
intoxication with CNF-1 results in diminished barrier
function with enhanced F-actin filament formation
[45]. Specific effects on tight junctional proteins have
not been reported. It is intriguing to note that in
intestinal epithelial cell monolayers both inactivation
and activation of Rho GTPases result in diminished
barrier function.
4.2. Toxins that destroy TJ structural elements
4.2.1. Bacteroides fragilis toxin (BFT), a zinc-
dependent metalloprotease toxin
BFT is encoded by a gene, bft, consisting of one
open reading frame of 1191 nucleotides that encodes a
protein of 397 amino acid residues [46,47]. Compar-
ison of the N-terminal sequence determined from
purified BFT with the predicted protein from the
nucleotide sequence suggests that BFT is synthesized
with three consecutive peptide domains: pre (or sig-
nal) sequence (18 amino acid residues), pro-peptide
(193 amino acid residues) and the mature protein (186
amino acid residues). This structure suggests that BFT
belong to the intramolecular chaperone protease fam-
ily [46]. Covalently linked pro-peptides in this family
(serving as the intramolecular chaperone) are essential
to both proper proteins folding for biologic activity
and secretion of the biologically active protein. Cur-
rently, no details are available on the intracellular
synthesis, processing and secretion of the BFT protein
by enterotoxigenic B. fragilis (ETBF) strains. Three
distinct chromosomal bft sequences (termed bft-1, bft-
2 and bft-3) have been reported that are 92–96%
identical in their predicted amino acid sequences with
the majority (>90%) of the amino acid differences
identified in the mature toxin protein [47]. BFT
proteins purified from culture supernatants of ETBF
strains secreting these distinct BFT isotypes exhibit
biochemical differences but only modest changes in
biologic activity to date.
BFT exhibits two major biologic activities, stimu-
lation of secretion in ligated ileal and colonic seg-
ments (lambs, calves, rabbits and rats) and alteration
of the morphology of epithelial cells (e.g. HT29/C1,
T84, Caco-2, MDCK) capable of forming tight junc-
tional complexes [i.e. ZO and zonula adherens (ZA)]
[48–50]. The cloned HT29/C1 cell line (a human
colonic carcinoma cell) has been studied most exten-
sively as a model for the mechanism of action of BFT.
These cells exhibit a rapid and striking change in
morphology when exposed to BFT without a loss of
viability [51]. Available data using inhibitors of endo-
somal function and intracellular vesicular trafficking
suggest that BFT is not internalized and, thus, is
thought to modify cell structure and function from
an extracellular location [50]. The half-maximal con-
centration of BFT-2 altering HT29/C1 cell morphol-
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807 801
ogy is f12.5 pM whereas it is f1 nM in polarized
T84 monolayers. Only 0.5 pM (0.01 ng/ml) BFT is
required to alter HT29/C1 cell morphology after an 18
h incubation. The potency of BFT in modifying
intestinal epithelial cell structure and function is
consistent with the hypothesis that ETBF (presumably
with release of small amounts of BFT) stimulate(s)
acute and possibly chronic intestinal pathology.
Intestinal tissues exhibit a submucosal inflamma-
tory response when infected with ETBF or treated
with purified BFT [52,53]. Higher doses of BFT lead
to secretion of mildly hemorrhagic fluid with patchy
mucosal wall hemorrhage. Consistent with these
observations, BFT stimulates intestinal epithelial cells
in vitro to secrete the polymorphonuclear cell chemo-
attractant, IL-8, in a dose-dependent manner [54,55].
It is unknown if BFT stimulates pro-inflammatory
chemokine secretion in vivo in animals or humans.
No pathology from human disease or studies to
evaluate intestinal inflammation in human ETBF
disease is yet available.
To date, all of the pathophysiologic outcomes
following BFT treatment of intestinal epithelial cells
have been linked to coincident morphologic changes
in these cells. When polarized monolayers of epithe-
lial cells (T84, MDCK, HT29, HT29/C1, Caco-2) are
treated with BFT in vitro, BFT decreases the resis-
tance of the epithelial monolayers in a dose- and
time-dependent manner [48,56]. Electron microscop-
ic analysis of T84 monolayers after BFT treatment
reveals swollen cells in which apical F-actin staining
is diminished and the microvilli are unraveled. Be-
tween some cells there is complete effacement of the
ZO (TJ) and the ZA [48]. However, increased F-
actin is detected at the basolateral pole of the
intestinal epithelial cells suggesting that BFT stim-
ulates a dynamic restructuring of cellular F-actin via
an as yet unknown mechanism. Consistent with this
hypothesis, total F-actin content of BFT-treated cells
is unchanged.
Recent data indicate that BFT acts as a protease
consistent with the predictions from its amino acid
sequence [50]. Of cellular structural proteins exam-
ined, only E-cadherin, the major structural protein of
the ZA, is cleaved by BFT in a time- and concentra-
tion-dependent manner [50]. Onset of E-cadherin
cleavage is detected within 1 min in HT29/C1 cells
preceding the first detected morphology changes
(detected at f10 min) and, similarly, re-synthesis of
E-cadherin correlates with recovery of HT29/C1 cell
morphology. Additional studies suggest that cleavage
of E-cadherin by BFT is a two step process in which
the extracellular domain of E-cadherin is first degrad-
ed in an ATP-independent manner (potentially directly
mediated by BFT) followed by the degradation of the
intracellular domain of E-cadherin in an ATP-depen-
dent manner (potentially mediated by one or more
cellular proteases). BFT was the first bacterial toxin
identified to remodel the intestinal epithelial cytoskel-
eton and F-actin architecture via cleavage of a cell
surface molecule.
4.2.2. Serine protease autotransporters of
enterobacteriaceae
A growing number of proteins have been found to
be secreted through the outer membrane of gram
negative bacteria in a mechanism similar to that first
described for the IgA protease of Neisseria [57].
Proteins secreted by this so-called autotransporter
mechanism carry a dedicated C-terminal domain,
which is thought to fold into a beta-barrel, through
which the mature protein exits the bacterium. A
family of these secreted proteins possesses a function-
al serine protease motif at a conserved position. The
proteases cleave a variety of substrates and are thought
to execute a variety of functions for their respective
pathogens [58].
One family of these proteases, called the serine
protease autotransporters of enterobacteriaceae (SPA-
TEs) are expressed by diarrheagenic and uropathogenic
E. coli and Shigella strains [59]. They are generally the
most abundant proteins in the supernatants of their host
strains when grown in laboratory conditions. Several
have been shown to induce cytopathic effects, but their
precise roles in pathogenesis have not been determined
for any of these proteins.
Many strains of enteroaggregative E. coli (EAEC)
carry the gene for a SPATE protein called Pet
(plasmid-encoded toxin) [60–62]. Navarro-Garcia et
al. showed that the Pet protein induced rounding of
HEp-2 and HT-29 cells in culture. Although actin
microfilaments were dissolved, no cleavage of actin
was detected. In vitro organ cultures of pediatric
colonic tissue revealed that EAEC strain 042 induced
less mucosal damage in the absence of an intact pet
gene. In addition, Pet was shown to elicit enterotoxic
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807802
effects on rat jejunal tissue mounted in Ussing
chamber.
Pet appears to act intracellularly. Studies using
confocal microscopy reveal internalization and traf-
ficking of the toxin in epithelial cells, and the effects
of Pet are blocked by preincubating cells with
Brefeldin A [60]. Recently, Sui et al. have repro-
duced Pet’s cellular effects by expressing the toxin
within the cytoplasm using mammalian expression
vectors [63].
Pet has been shown to cleave fodrin, a component
of the membrane cytoskeleton [64], and disruption of
fodrin precedes dissolution of actin stress fibers (R.
Cappello and J. Nataro, unpublished). Fodrin is
thought to assist in maintenance of membrane
domains, in providing stability and shape to organ-
elles and in linking the membrane to transport proteins
and to the major filament systems [65,66]. Disruption
of the fodrin system could thus induce a myriad of
cellular abnormalities, including disorders of barrier
function and ion secretion.
4.2.3. Vibrio cholerae hemagglutinin protease
The hemagglutinin protease (HA/P) elaborated by
V. cholerae represents a typical example of direct
effects of a bacterial protease on TJ proteins. HA/P, a
toxin belonging to the family of bacterial metallopro-
teases, possesses multiple functions, including the
activation of cholera toxin (CT). Interestingly, HA/P
also inactivates the CTXf that houses the V. cholerae
enterotoxin CT, Zonula occluden toxin (Zot), and
Accessory cholera enterotoxin (Ace) (see below).
Studies by Wu and collaborators showed that epithe-
lial cell monolayers exposed to HA/P showed de-
creased TER and disruption of ZO-1 and the actin
cytoskeleton in a dose-dependent manner [67]. The
same authors have recently shown that HA/P also
degrades occludin in a dose- and time-dependent
fashion [68]. Based on the sizes of the degradation
products, the hydrophilicity plot of occludin, and the
specificity of the antibodies used, it was predicted that
HA/P cleaves occludin in two fragments, one repre-
senting the cytosolic COOH-terminus plus the adja-
cent membrane domain (f35 kDa), and the second
fragment (f50 kDa) being composed by the remain-
ing occludin molecule [68]. The HA/P cleaving
activity was prevented by bacterial metalloprotease
inhibition, suggesting that the TJ disruption caused by
the toxin can be related to either its intrinsic metal-
loprotease activity or to the activation by HA/P of an
endogenous metalloprotease.
4.3. Toxins that affect TJ competency by stimulating
host signaling events
4.3.1. Zonula occludens toxin
Several microorganisms have been shown to exert
their effect on intercellular TJ complex by activating
host cellular signal transduction pathways [69]. Con-
versely, functional mimicry of an endogenous modu-
lator affecting TJ permeability has been proposed for
the Zot elaborated by V. cholerae [70,71]. Zot is a
single polypeptide chain of 44.8 kDa encoded by the
bacteriophage CTXf present in toxigenic strains of V.
cholerae [72]. Zot possesses multiple domains that
suggest a dual function of the protein as a morpho-
genetic phage peptide for the V. cholerae phage
CTXf and as an enterotoxin that modulates intestinal
TJ [73]. Zot localizes in the bacterial outer membrane
of V. cholerae with subsequent cleavage and secretion
of a C-terminal fragment in the host intestinal milieu
[73]. Structure–function analysis of the toxin sug-
gested that these two fragments have distinctive
biological functions [74]. Its f33 kDa N-terminal
portion possesses homology with pI proteins of other
filamentous bacteriophages [73] and is, therefore,
possibly involved in the CTXf phage assembly, while
the f12 kDa C-terminal fragment of the toxin seems
to be responsible for the permeating action on intes-
tinal TJ [74]. Interestingly, the Zot C-terminal frag-
ment shares functional analogies with zonulin, the
recently described Zot mammalian analogue involved
in TJ modulation [75]. Amino acid comparison be-
tween the Zot active fragment and zonulin, combined
with site-directed mutagenesis experiments, con-
firmed the presence of an octapeptide receptor-bind-
ing domain toward the N-terminus of the processed
Zot [74].
Zot effects on TJ modulation are mediated by a
cascade of intracellular events that lead to a PKCa-
dependent polymerization of actin microfilaments
strategically localized to regulate the paracellular
pathway [76] (Fig. 2). The toxin exerts this effect by
interacting with the zonulin surface receptor, whose
distribution varies within the intestine. The zonulin
receptor is detectable in the jejunum and distal ileum,
Fig. 2. Proposed Zot intracellular signaling leading to the opening of intestinal TJ. Zot interacts with a specific surface receptor (1) whose
distribution within the intestine varies. The protein is then internalized and activates phospholipase C (2) that hydrolyzes phosphatidyl inositol
(3) to release inositol 1,4,5-tris phosphate (PPI-3) and diacylglycerol (DAG) (4). PKCa is then activated (5), either directly (via DAG) (4) or
through the release of intracellular Ca2 + (via PPI-3) (4a). Membrane-associated, activated PKCa (6) catalyzes the phosphorylation of target
protein(s), with subsequent polymerization of soluble G-actin in F-actin (7). This polymerization causes the rearrangement of the filaments of
actin and the subsequent displacement of proteins (including ZO-1) from the junctional complex (8). As a result, intestinal TJ become looser.
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807 803
but not in the colon, and decreases along the villous–
crypt axis [77]. This receptor distribution coincides
with the regional effect of Zot on intestinal perme-
ability [77] and with the preferential F-actin redistri-
bution induced by Zot in the mature cells of the villi
[76]. These data also suggest that the expression of the
zonulin receptor(s) is up-regulated during enterocyte
differentiation. This hypothesis is supported by the
observation that human intestinal epithelial CaCo2cells (which resemble the mature absorptive enteric
cell of the villi), but not crypt-like T84 cells, express
this receptor(s) on their surface [78]. The paucity of
Zot binding in the crypt area may also reflect the fact
that this region is already leaky as compared to the
more mature epithelium of the tip of the villi [79], and
thus might not need to express a significant amount of
the zonulin receptor involved in TJ regulation.
Following binding to the zonulin receptor, Zot
induces actin polymerization [76], followed by ZO-
1–occludin and ZO-1–claudin disengagement [80]
and down-regulation of occludin gene expression
[81]. These changes occur as soon as 15–30 min
following Zot exposure both in enterocyte cell lines
and whole intestinal tissues and are temporally coin-
cident with TJ disassembly [80]. However, the de-
creased occludin gene expression is followed by
down-regulation of the protein pool only 6–9 h post-
Zot exposure (A. Fasano, unpublished), suggesting a
two-step process in which the early ZO-1–occludin
disengagement (i.e. rapid and reversible TJ disassem-
bly) is followed by a more prolonged effect on TJ
(related to decreased occludin protein pool) if enteric
cells are chronically exposed to Zot. Taken together,
these data suggest that Zot regulates TJ in a rapid,
reversible, reproducible fashion, and activates intra-
cellular signals involved in zonulin-mediated modu-
lation of the paracellular pathway.
Recent data seems to suggest that the Zot-activat-
ed zonulin system plays an important role in gut
innate immunity [82]. In the absence of enteric
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807804
infections, the mammalian small intestine is virtually
sterile. The colonization of the proximal gut by
enteric microorganisms (even without apparent mu-
cosal damage or elaboration of specific toxins) may
promote a leaky intestine [83], however, the mech-
anism(s) by which proximal bacterial contamination
disturbs intestinal TJ permeability remains unclear. It
has been recently reported that both normal enteric
bacterial flora and pathogenic bacteria each induce
alteration of TJ competency as suggested by changes
in epithelial resistance and increased passage of
inulin. These changes were mirrored by the concom-
itant luminal secretion of zonulin in organ culture
systems and occurred even when nonviable bacteria
were introduced [82]. That interaction of bacteria
with the intestinal mucosa induces zonulin release,
irrespective of their pathogenic traits or viability, and
can be interpreted as a bacteria-independent host
defense mechanism (innate immunity) that reacts to
the abnormal presence of microorganisms on the
surface of the small intestine. Following the zonu-
lin-induced opening of TJ, water is secreted into the
intestinal lumen following hydrostatic pressure gra-
dients [77] and bacteria are ‘‘flushed out’’ from the
small intestine.
4.3.2. C. perfringens enterotoxin (CPE)
Native CPE is a 35 kDa peptide that acts as a
potent cytotoxin in in vitro and in vivo studies
(reviewed in Ref. [84]). Detailed studies suggest that
CPE binds irreversibly with several membrane pro-
teins yielding a pore-forming complex resulting in
rapid cell death. In rabbits, intestinal secretion is
always associated with histopathologic damage. Re-
cently, a carboxy-terminal fragment of CPE has been
shown to cleave claudins 3 and 4, key proteins of the
ZO, resulting in diminished barrier function without
cytotoxicity [85,86]. These data provided the first
evidence indicating the physiologic importance of
the claudin proteins in the barrier function of the
ZO. Whether this mechanism contributes to the in
vivo effects of CPE is unknown.
4.3.3. V. cholerae RTX (‘repeats-in-toxin’) toxin
The RTX toxin (encoded by rtxA) produced by El
Tor and O139, but not by classical, V. cholerae
strains elicits rounding of both epithelial and non-
epithelial cell lines [87]. This cellular phenotype
occurs by an unknown mechanism that involves
depolymerization of F-actin and cross-linking of G-
actin into dimers, trimers and higher multimers [88].
Initial experiments examining the effect of mutant El
Tor strains on the physiology of human colonic
epithelial cell (T84) monolayers revealed that pro-
duction of the RTX toxin was associated with a loss
of barrier function consistent with its described in
vitro effects on F- and G-actin [89].
5. Concluding remarks and future directions
The paracellular pathway was once considered to
be exclusively the route for passive, unregulated
passage of water, electrolytes, and small molecules.
Its contribution to transepithelial transports was,
therefore, judged to be simply secondary to the
active, transcellular transport processes. It is now
becoming apparent that the elements that govern this
pathway; i.e. the TJ, are extremely dynamic struc-
tures involved in developmental, physiological, and
pathological circumstances. An increased number of
toxins had been shown to exert their pathogenic
action by altering TJ competency, either directly or
through cytoskeletal changes. The field lies at the
nexus of bacteriology, protein chemistry and cell
biology, and provides a prominent example of syn-
ergistic research among scientific disciplines. The
remarkable examples of toxins targeting intercellular
TJ described in this chapter are paving the way to
new knowledge that will most likely lead to a better
understanding of the regulation of intercellular TJ in
health and disease, and will offer innovative strate-
gies to deliver drugs and vaccines in a more efficient
and non-invasive fashion.
References
[1] A.S. Fanning, L.L. Mitic, J.M. Anderson, Transmembrane
proteins in the tight junction barrier, J. Am. Soc. Nephrol.
10 (1999) 1337–1345.
[2] J.L. Madara, K. Dharmsathaphorn, Occluding junction struc-
ture– function relationships in a cultured epithelial monolayer,
J. Cell Biol. 101 (1985) 2124–2133.
[3] M. Furuse, T. Hirase, M. Itoh, A. Nagafuchi, S. Yonemura, S.
Tsukita, Occludin: a novel integral membrane protein localiz-
ing at tight junctions, J. Cell Biol. 123 (1993) 1777–1788.
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807 805
[4] M. Saitou, Y. Ando-Akatsuka, M. Itoh, M. Furuse, J. Inazawa,
K. Fujimoto, S. Tsukita, Mammalian occludin in epithelial
cells: its expression and subcellular distribution, Eur. J. Cell
Biol. 73 (1997) 222–231.
[5] M. Furuse, K. Fujita, T. Hiiragi, K. Fujimoto, S. Tsukita,
Claudin-1 and -2: novel integral membrane proteins localizing
at tight junctions with no sequence similarity to occludin,
J. Cell Biol. 141 (1998) 1539–1550.
[6] C. Rahner, L.L. Mitic, J.M. Anderson, Heterogeneity in
expression and subcellular localization of claudins 2, 3, 4,
and 5 in the rat liver, pancreas, and gut, Gastroenterology
120 (2001) 411–422.
[7] M. Furuse, H. Sasaki, S. Tsukita, Manner of interaction of
heterogeneous claudin species within and between tight junc-
tion strands, J. Cell Biol. 147 (1999) 891–903.
[8] J. Katahira, N. Inoue, Y. Horiguchi, M. Matsuda, N. Sugimoto,
Molecular cloning and functional characterization of the recep-
tor for Clostridium perfringens enterotoxin, J. Cell Biol. 136
(1997) 1239–1247.
[9] J. Katahira, H. Sugiyama, N. Inoue, Y. Horiguchi, M. Matsu-
da, N. Sugimoto, Clostridium perfringens enterotoxin utilizes
two structurally related membrane proteins as functional re-
ceptors in vivo, J. Biol. Chem. 272 (1997) 26652–26658.
[10] I. Martin-Padura, S. Lostaglio, M. Schneemann, L. Williams,
M. Romano, P. Fruscella, C. Panzeri, A. Stoppacciaro, L.
Ruco, A. Villa, D. Simmons, E. Dejana, Junctional adhesion
molecule, a novel member of the immunoglobulin superfam-
ily that distributes at intercellular junctions and modulates
monocyte transmigration, J. Cell Biol. 141 (1998) 117–127.
[11] B.R. Stevenson, J.D. Siliciano, M.S. Mooseker, D.A.
Goodenough, Identification of ZO-1: a high molecular
weight polypeptide associated with the tight junction (zon-
ula occludens) in a variety of epithelia, J. Cell Biol. 103
(1986) 755–766.
[12] T. Tsukamoto, S.K. Nigam, Tight junction proteins form large
complexes and associate with the cytoskeleton in an ATP
depletion model for reversible junction assembly, J. Biol.
Chem. 272 (1997) 16133–16139.
[13] B. Gumbiner, T. Lowenkopf, D. Apatira, Identification of a
160-kDa polypeptide that binds to the tight junction protein
ZO-1, Proc. Natl. Acad. Sci. USA 88 (1991) 3460–3464.
[14] J. Haskins, L. Gu, E.S. Wittchen, J. Hibbard, B.R. Stevenson,
ZO-3, a novel member of the MAGUK protein family found
at the tight junction, interacts with ZO-1 and occludin, J. Cell
Biol. 141 (1998) 199–208.
[15] M.S. Balda, K. Matter, Tight junctions, J. Cell Sci. 111 (1998)
541–547.
[16] B.H. Keon, S. Schafer, C. Kuhn, C. Grund, W.W. Franke,
Symplekin, a novel type of tight junction plaque protein, J.
Cell Biol. 134 (1996) 1003–1018.
[17] C.J. Gottardi, M. Arpin, A.S. Fanning, D. Louvard, The
junction-associated protein, zonula occludens-1, localizes to
the nucleus before the maturation and during the remodeling
of cell –cell contacts, Proc. Natl. Acad. Sci. USA 93 (1996)
10779–10784.
[18] B. Gumbiner, Structure, biochemistry, and assembly of epithe-
lial tight junctions, Am. J. Physiol. 253 (1987) C749–C758.
[19] J.L. Madara, D. Barenberg, S. Carlson, Effects of cytochalasin
D on occluding junctions of intestinal absorptive cells: further
evidence that the cytoskeleton may influence paracellular per-
meability and junctional charge selectivity, J. Cell Biol. 102
(1986) 2125–2136.
[20] D. Drenchahn, R. Dermietzel, Organization of the actin fila-
ment cytoskeleton in the intestinal brush border: a quantita-
tive and qualitative immunoelectron microscope study,
J. Cell Biol. 107 (1988) 1037–1048.
[21] F. Hecht, Expression of the catalytic domain of myosin light
chain kinase increases paracellular permeability, Am. J. Phys-
iol. 271 (1996) C1678–C1684.
[22] K. Tsuneo, U. Brauneis, Z. Gatmaitan, I. Arias, Extracellular
ATP, intracellular calcium and canalicular contraction in rat
hepatocye doublets, Hepatology 14 (1991) 640–647.
[23] A. Fasano, C. Fiorentini, G. Donelli, S. Uzzau, J.B. Kaper,
K. Margaretten, X. Ding, S. Guandalini, L. Comstock, S.E.
Goldblum, Zonula occludens toxin modulates tight junctions
through protein kinase C-dependent actin reorganization, in
vitro, J. Clin. Invest. 96 (1995) 710–720.
[24] L.L. Mitic, C.M. Van Itallie, Occludin and claudins: trans-
membrane proteins of the tight junction, Tight Junctions,
CRC Press, Boca Raton, FL, 2001, pp. 213–230.
[25] M. Itoh, M. Furuse, K. Morita, K. Kubota, M. Saitou, S.
Tsukita, Direct binding if three tight junction-associated
MAGUKs, ZO-1, ZO-2, and ZO-3, with the COOH termini
of claudins, J. Cell Biol. 147 (1999) 1351–1363.
[26] G. Schmidt, K. Aktories, Bacterial cytotoxins target Rho
GTPases, Naturwissenschaften 85 (1998) 253–261.
[27] G. Schmidt, K. Aktories, Rho GTPase-activating toxins:
cytotoxic necrotizing factors and dermonecrotic toxin,
Methods Enzymol. 325 (2000) 125–136.
[28] H.J. Schnittler, S.W. Schneider, H. Raifer, F. Luo, P. Dieterich,
I. Just, K. Aktories, Role of actin filaments in endothelial
cell – cell adhesion and membrane stability under fluid shear
stress, Pflug. Arch. 442 (2001) 675–687.
[29] I. Just, F. Hofmann, K. Aktories, Molecular mode of action
of the large clostridial cytotoxins, Curr. Top. Microbiol.
Immunol. 250 (2000) 55–83.
[30] I. Just, F. Hofmann, H. Genth, R. Gerhard, Bacterial protein
toxins inhibiting low-molecular-mass GTP-binding proteins,
Int. J. Med. Microbiol. 291 (2001) 243–250.
[31] S.P. Borriello, Pathogenesis of Clostridium difficile infection,
J. Antimicrob. Chemother. 41 (1998) 13–19.
[32] K. Aktories, Bacterial toxins that target Rho proteins, J. Clin.
Invest. 99 (1997) 827–829.
[33] S. Narumija, T. Ishizaki, N. Watanabe, Rho effectors and
reorganization of actin cytoskeleton, FEBS Lett. 410 (1997)
68–72.
[34] C. Pothoulakis, Pathogenesis of Clostridium difficile-asso-
ciated diarrhoea, Eur. J. Gastroenterol. Hepatol. 8 (1996)
1041–1047.
[35] C. Pothoulakis, Effects of Clostridium difficile toxins on epi-
thelial cell barrier, Ann. New York Acad. Sci. 915 (2000)
347–356.
[36] C. Pothoulakis, J.T. Lamont, Microbes and microbial toxins:
paradigms for microbial–mucosal interactions: II. The inte-
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807806
grated response of the intestine to Clostridium difficile tox-
ins, Am. J. Physiol. Gastrointest. Liver Physiol. 280 (2001)
G178–G183.
[37] D. He, S.J. Hagen, C. Pothoulakis, M. Chen, N.D. Medina,
M. Warny, J.T. LaMont, Clostridium difficile toxin A causes
early damage to mitochondria in cultured cells, Gastroente-
rology 119 (2000) 139–150.
[38] C. Alcantara, W.F. Stenson, T.S. Steiner, R.L. Guerrant, Role
of inducible cyclooxygenase and prostaglandins in Clostridium
difficile toxin A-induced secretion and inflammation in an ani-
mal model, J. Infect. Dis. 184 (2001) 648–652.
[39] J.F. Richard, L. Petit, M. Gibert, J.C. Marvaud, C. Bouchaud,
M.R. Popoff, Bacterial toxins modifying the actin cytoskeleton,
Int. Microbiol. 2 (1999) 185–194.
[40] C. Wilde, K. Aktories, The Rho-ADP-ribosylating C3 exoen-
zyme from Clostridium botulinum and related C3-like trans-
ferases, Toxicon 39 (2001) 1647–1660.
[41] A. Nusrat, M. Giry, J.R. Turner, S.P. Colgan, C.A. Parkos,
D. Carnes, E. Lemichez, P. Bouquest, J.L. Madara, Rho
protein regulates tight junctions and perijunctional actin
organization in polarized epithelia, Proc. Natl. Acad. Sci.
USA 92 (1995) 10629–10635.
[42] A. Andreu, A.E. Stapleton, C. Fennell, H.A. Lockman, M.
Xercavins, F. Fernandez, W.E. Stamm, Urovirulence determi-
nants in Escherichia coli strains causing prostatitis, J. Infect.
Dis. 176 (1997) 464–469.
[43] P. Boquet, The cytotoxic necrotizing factor 1 (CNF-1) from
Escherichia coli, Toxicon 39 (2001) 1673–1680.
[44] K.E. Rippere-Lampe, A.D. O’Brien, R. Conran, H.A. Lock-
man, Mutation of the gene encoding cytotoxic necrotizing fac-
tor type 1 (cnf(1)) attenuates the virulence of uropathogenic
Escherichia coli, Infect. Immun. 69 (2001) 3954–3964.
[45] R. Gerhard, G. Schmidt, F. Hofmann, K. Aktories, Activa-
tion of Rho GTPases by Escherichia coli cytotoxic necro-
tizing factor 1 increases intestinal permeability in Caco-2
cells, Infect. Immun. 66 (1998) 5125–5131.
[46] C.L. Sears, The toxins of Bacteroides fragilis, Toxicon 39
(2001) 1737–1746.
[47] S. Wu, L.A. Dreyfus, A.O. Tzianabos, C. Hayashi, C.L.
Sears, Diversity of the metalloprotease toxin produced by
enterotoxigenic Bacteroides fragilis, Infect. Immun. 70
(2002) 2463–2471.
[48] F.G. Chambers, S.S. Koshy, R.F. Saidi, D.P. Clark, R.D.
Moore, C.L. Sears, Bacteroides fragilis toxin exhibits polar
activity on monolayers of human intestinal epithelial cells
(T84 cells) in vitro, Infect. Immun. 65 (1997) 3561–3570.
[49] R.J. Obiso Jr., D.M. Lyerly, R.L. Van Tassell, T.D. Wilkins,
Proteolytic activity of the Bacteroides fragilis enterotoxin
causes fluid secretion and intestinal damage in vivo, Infect.
Immun. 63 (1995) 3820–3826.
[50] S. Wu, K.C. Lim, J. Huang, R.F. Saidi, C.L. Sears, Bac-
teroides fragilis enterotoxin cleaves the zonula adherens
protein, E-cadherin, Proc. Natl. Acad. Sci. USA 95
(1998) 14979–14984.
[51] R.F. Saidi, C.L. Sears, Bacteroides fragilis toxin rapidly
intoxicates human intestinal epithelial cells (HT29/C1) in
vitro, Infect. Immun. 64 (1996) 5029–5034.
[52] C.L. Sears, The toxins of Bacteroides fragilis, Toxicon 39
(2001) 1737–1746.
[53] C.L. Sears, L.L. Myers, A. Lazenby, R.L. Van Tassell, Entero-
toxigenic Bacteroides fragilis, Clin. Infect. Dis. 20 (Suppl. 2)
(1995) S142–S148.
[54] L. Sanfilippo, C.K. Li, R. Seth, T.J. Balwin, M.G. Menozzi,
Y.R. Mahida, Bacteroides fragilis enterotoxin induces the
expression of IL-8 and transforming growth-factor-beta
(TGF-beta) by human colonic epithelial cells, Clin. Exp.
Immunol. 119 (2000) 456–463.
[55] J.M. Kim, Y.K. Oh, Y.J. Kim, H.B. Oh, Y.J. Cho, Polarized
secretion of CXC chemokines by human intestinal epithelial
cells in response to Bacteroides fragilis enterotoxin: NF-kappa
B plays a major role in the regulation of IL-8 expression, Clin.
Exp. Immunol. 123 (2001) 421–427.
[56] R.J. Obiso Jr., A.O. Azghani, T.D. Wilkins, The Bacteroides
fragilis toxin fragilysin disrupts the paracellular barrier of
epithelial cells, Infect. Immun. 65 (1997) 1431–1439.
[57] I.R. Henderson, F. Navarro-Garcia, J.P. Nataro, The great
escape: structure and function of the autotransporter proteins,
Trends Microbiol. 6 (1998) 370–378.
[58] I.R. Henderson, J.P. Nataro, Virulence functions of autotrans-
porter proteins, Infect. Immun. 69 (2001) 1231–1243.
[59] P.R. Dutta, R. Cappello, F. Navarro-Garcia, J.P. Nataro,
Functional comparison of serine protease autotransporters
of enterobacteriaceae, Infect. Immun. 70 (2002) 7105–7113.
[60] F. Navarro-Garcia, A. Canizalez-Roman, J. Luna, C. Sears, J.P.
Nataro, Plasmid-encoded toxin of enteroaggregative Escheri-
chia coli is internalized by epithelial cells, Infect. Immun. 69
(2001) 1053–1060.
[61] F. Navarro-Garcia, C. Eslava, J.M. Villaseca, R. Lopez-Revilla,
J.R. Czeczulin, S. Srinivas, J.P. Nataro, A. Cravioto, In vitro
effects of a high-molecular-weight heat-labile enterotoxin from
enteroaggregative Escherichia coli, Infect. Immun. 66 (1998)
3149–3154.
[62] F. Navarro-Garcia, C. Sears, C. Eslava, A. Cravioto, J.P. Nataro,
Cytoskeletal effects induced by pet, the serine protease ente-
rotoxin of enteroaggregative Escherichia coli, Infect. Immun.
67 (1999) 2184–2192.
[63] B.Q. Sui, P.R. Dutta, J.P. Nataro, Intracellular expression of
the plasmid-encoded toxin from enteroaggregative Escheri-
chia coli, Infect. Immun. 71 (2003) 5364–5370.
[64] J.M. Villaseca, F. Navarro-Garcia, G.Mendoza-Hernandez, J.P.
Nataro, A. Cravioto, C. Eslava, Pet toxin from enteroaggrega-
tive Escherichia coli produces cellular damage associated with
fodrin disruption, Infect. Immun. 68 (2000) 5920–5927.
[65] M.A. De Matteis, J.S. Morrow, The role of ankyrin and fodrin
in membrane transport and domain formation, Curr. Opin. Cell
Biol. 10 (1998) 542–549.
[66] M.A. De Matteis, J.S. Morrow, Fodrin tethers and mesh in the
biosynthetic pathway, J. Cell Sci. 113 (2000) 2331–2343.
[67] Z. Wu, D. Milton, P. Nybom, A. Sjo, K.E. Magnusson, Vibrio
cholerae hemagglutinin/protease (HA/protease) causes mor-
phological changes in cultured epithelial cells and perturbs
their paracellular barrier function, Microb. Pathog. 21 (1996)
111–123.
[68] Z. Wu, P. Nybom, K.E. Magnusson, Distinct effects of Vibrio
A. Fasano, J.P. Nataro / Advanced Drug Delivery Reviews 56 (2004) 795–807 807
cholerae haemagglutinin/protease on the structure and locali-
zation of the tight junction-associated proteins occludin and
ZO-1, Cell Microbiol. 2 (2000) 11–17.
[69] A. Fasano, Cellular microbiology: can we learn cell physio-
logy from microorganisms? Am. J. Physiol. 276 (1999)
C765–C776.
[70] A. Fasano, B. Baudry, D.W. Pumplin, S.S. Wasserman, B.D.
Tall, J.M. Ketley, J.B. Kaper, Vibrio cholerae produces a sec-
ond enterotoxin, which affects intestinal tight junctions, Proc.
Natl. Acad. Sci. USA 88 (1991) 5242–5246.
[71] B. Baudry, A. Fasano, J.M. Ketley, J.B. Kaper, Cloning of a
gene (ZOT) encoding a new toxin produced by Vibrio cholerae,
Infect. Immun. 60 (1991) 428–434.
[72] M.K.Waldor, J.J. Mekalanos, Science 272 (1996) 1910–1914.
[73] S. Uzzau, P. Cappuccinelli, A. Fasano, Expression of Vibrio
cholerae zonula occludens toxin and analysis of its subcellular
localization, Microb. Pathog. 27 (1999) 377–385.
[74] M. Di Pierro, R. Lu, S. Uzzau, W. Wang, K. Margaretten, C.
Pazzani, F. Maimone, A. Fasano, Zonula occludens toxin
structure– function analysis: identification of the fragment bi-
ologically active on tight junctions and of the zonulin receptor
binding domain, J. Biol. Chem. 276 (2001) 19160–19165.
[75] W. Wang, S. Uzzau, S.E. Goldblum, A. Fasano, Human
zonulin, a potential modulator of intestinal tight junctions,
J. Cell Sci. 113 (2000) 4435–4440.
[76] A. Fasano, C. Fiorentini, G. Donelli, S. Uzzau, J.B. Kaper,
K. Margaretten, X. Ding, S. Guandalini, L. Comstock, S.E.
Goldblum, Zonula occludens toxin modulates tight junctions
through protein kinase C-dependent actin reorganization, in
vitro, J. Clin. Invest. 96 (1995) 710–720.
[77] A. Fasano, S. Uzzau, C. Fiore, K. Margaretten, The entero-
toxic effect of zonula occludens toxin (Zot) on rabbit small
intestine involves the paracellular pathway, Gastroenterology
112 (1997) 839–846.
[78] S. Uzzau, R. Lu, W. Wang, C. Fiore, A. Fasano, Purification
and preliminary characterization of the zonula occludens toxin
receptor from human (CaCo2) and murine (IEC6) intestinal
cell lines, FEMS Microbiol. Lett. 194 (2001) 1–5.
[79] M.A. Marcial, S.L. Carlson, J.L. Madara, Partitioning of
paracellular conductance along the ileal crypt-villus axis: a
hypothesis based on structural analysis with detailed consi-
deration of tight junction structure– function relationships,
J. Membr. Biol. 80 (1984) 59–70.
[80] T. Watts, T. Kiser, R. Macatangay, S. Goldblum, A. Fasano,
Zonula occludens toxin (Zot) modulates disassembly of intes-
tinal tight junctions by altering the association between ZO-1
and occludin, Pediatr. Res. 49 (2001) 119A.
[81] M.R. Di Pierro, S. Drago, M. Thakar, R. Lu, F. Maimone, A.
Fasano, Zonulin occludens toxin induces a decreased expre-
ssion of the tight junction protein occludin, Gastroenterology
122 (2002) A405.
[82] R. El Asmar, P. Panigrahi, P. Bamford, I. Berti, T. Not, G.V.
Coppa, C. Catassi, A. Fasano, Host-dependent activation of
the zonulin system is involved in the impairment of the gut
barrier function following bacterial colonization, Gastroente-
rology 123 (2002) 1607–1615.
[83] A. Fasano, Pathological and therapeutical implications of
macromolecule passage through the tight junction, Tight Junc-
tions, CRC Press, Boca Raton, FL, 2001, pp. 697–722.
[84] M.R. Sarker, U. Singh, B.A. McClane, An update on Clos-
tridium perfringens enterotoxin, J. Nat. Toxins 9 (2000)
251–266.
[85] B.A. McClane, Clostridium perfringens enterotoxin and intes-
tinal tight junctions, Trends Microbiol. 8 (2000) 145–146.
[86] B.A. McClane, The complex interactions between Clostridium
perfringens enterotoxin and epithelial tight junctions, Toxicon
39 (2001) 1781–1791.
[87] W. Lin, K.J. Fullner, R. Clayton, J.A. Sexton, M.B. Rogers,
K.E. Calia, S.B. Calderwood, C. Fraser, J.J. Mekalanos, Iden-
tification of a Vibrio cholerae RTX toxin gene cluster that is
tightly linked to the cholera toxin prophage, Proc. Natl. Acad.
Sci. USA 96 (1999) 1071–1076.
[88] K.J. Fullner, J.J. Mekalanos, In vivo covalent cross-linking of
cellular actin by the Vibrio cholerae RTX toxin, EMBO J. 19
(2000) 5315–5323.
[89] K.J. Fullner, W.I. Lencer, J.J. Mekalanos, Vibrio cholerae-
induced cellular responses of polarized T84 intestinal epithe-
lial cells are dependent on production of cholera toxin and the
RTX toxin, Infect. Immun. 69 (2001) 6310–6317.