Phage display and Shiga toxin neutralizers

Accepted Manuscript Phage display and Shiga toxin neutralizers Bernedo-Navarro, Robert Alvin, Tomomasa Yano

PII:

S0041-0101(16)30027-7

DOI:

10.1016/j.toxicon.2016.02.009

Reference:

TOXCON 5309

To appear in:

Toxicon

Received Date: 4 June 2015 Revised Date:

3 February 2016

Accepted Date: 11 February 2016

Please cite this article as: Bernedo-Navarro, Alvin, R., Yano, T., Phage display and Shiga toxin neutralizers, Toxicon (2016), doi: 10.1016/j.toxicon.2016.02.009. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Mini Review

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Title: Phage display and Shiga toxin neutralizers

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Author: Bernedo-Navarro, Robert Alvin1 and Tomomasa Yano1

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(UNICAMP) – SP- Brazil

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E-mail: [email protected]

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Corresponding author: Bernedo-Navarro, Robert Alvin

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Department of Genetics, Evolution and Bioagents – University of Campinas

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Keywords: Escherichia coli; STEC; Shiga toxins; Phage display

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Abstract

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The current work presents an overview of the use of phage display technology for the

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identification and characterization of potential neutralizing agents for Shiga toxins. The

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last major Shiga toxin-associated disease outbreak, which took place in Germany in 2011,

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showed the international community that Shiga toxins remain a serious threat to public

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health. This is also demonstrated by the lack of specific therapies against Shiga toxin-

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induced Hemolytic Uremic Syndrome (HUS). Since its inception, phage display

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technology has played a key role in the development of antigen-specific (poly)-peptides

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or antibody fragments with specific biological properties. Herein, we review the current

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literature regarding the application of phage display to identify novel neutralizing agents

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against Shiga toxins. We also briefly highlight reported discoveries of peptides and heavy

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chain antibodies (VHH fragments or nanobodies) that can neutralize the cellular damage

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caused by these potent toxins.

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Pathogen

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Shiga toxin-producing Escherichia coli (STEC) are a heterogeneous and potentially fatal

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group of microorganisms that produce potent cytotoxins called Shiga toxins (Stxs). These

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toxins are similar to those produced by Shigella dysenteriae type 1.

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In addition to asymptomatic infections, STEC can cause the following clinical

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manifestations: acute watery diarrhea; bloody diarrhea; hemorrhagic colitis; hemolytic

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uremic syndrome (HUS), a life-threatening thrombotic microangiopathy leading to acute

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renal dysfunction approximately one week after onset of diarrhea; and death (Pennington,

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2010).

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E. coli serotype O157:H7 is the most common member of the STEC pathotype and the

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leading cause of pediatric HUS (Banatvala et al., 2001; Verweyen et al., 1999). However,

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over 400 non-O157 serotypes isolated from different sources have reported involvement

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in human disease (Bettelheim, 2007; Mora et al., 2011).

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It has previously been reported that children, the elderly and immunocompromised

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individuals are generally more susceptible to STEC infections; however, a recent E. coli

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O104 outbreak in Germany showed that infected healthy adults can also present with

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severe complications.

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The Shiga toxins encoded by the stx1 and stx2 genes, which are carried by lysogenic

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lambdoid phages, are the main virulence factors associated with STEC. Strains carrying

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the stx2 gene (mainly the stx2EDL933 subtype) are not only potentially more virulent but

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also are more frequently related to HUS than those harboring only stx1 and those carrying

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both genes (Friedrich et al., 2002; Schmidt et al., 1995).

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Among the STEC strains, seven serogroups have been more frequently associated with

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severe human illnesses, such as hemorrhagic colitis (HC) and HUS. These strains are also

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referred as enterohemorrhagic E. coli (EHEC) (Delannoy et al., 2013; Levine, 1987;

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Nataro and Kaper, 1998). In developed countries in the Northern Hemisphere, serotypes

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O26:H11, O45:H2, O103:H2, O111:H8, O121:H19, O145:H28, and O157:H7, as well as

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their non-motile derivatives, are considered the seven “priority” STEC serotypes (also

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referred as the “top 7” EHEC serotypes). These seven EHEC serotypes can be subdivided

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into seropathotypes (SPTs) A and B based on their phenotypic and molecular

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characteristics and the clinical features of their associated diseases. There are a total of

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five SPTs, which are denoted A, B, C, D, and E according to decreasing rank of

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pathogenicity (Karmali et al., 2003).

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In addition to Stx production, the top 7 serotypes harbor the locus of enterocyte

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effacement (LEE), a genomic island encoding intimin, which participates in bacterial

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colonization of the gut and in attaching-and-effacing (A/E) lesions of the intestinal

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mucosa (Nataro and Kaper, 1998), in addition to regulatory elements, a type III secretion

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system, and secreted effector proteins, as well as their cognate chaperones (Elliott et al.,

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1998; Perna et al., 1998).

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Other reported virulence-associated markers include a plasmid-encoded enterohemolysin

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and, in strains lacking eae, an autoagglutinating adhesin (Saa) potentially involved in

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adhesion (Paton et al., 2001). This plasmid is present in STEC O157 and in non-O157

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strains. Furthermore, STEC strains belonging to identical pulsed-field gel electrophoresis

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types can be further discriminated based on their plasmid-encoded genes, including

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hemolysin (ehxA), a catalase-peroxidase (katP), an extracellular serine protease (espP), a

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zinc metalloprotease (stcE, also called tagA), and a subtilase cytotoxin (subAB), among

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others (Etcheverría and Padola, 2013).

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STEC strains are innocuous to ruminants and are transient members of their

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gastrointestinal flora. Cattle are the main STEC reservoir implicated in human disease

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(Nastasijevic et al., 2008), and STEC strains have also been found in the gastrointestinal

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tracts of sheep, pigs, goats, dogs, and cats (Paton and Paton, 1998). Fecal contamination

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of meat during slaughter is the most common route for transmission to humans (Chase-

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Topping et al., 2008; Kudva et al., 1999). In this way, foods of bovine origin, especially

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undercooked ground beef and unpasteurized milk, constitute important sources of human

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infection (Griffin and Tauxe, 1991; Rangel et al., 2005). Other sources linked to sporadic

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infection and outbreaks of illness in humans include lettuce, alfalfa sprouts, radish

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sprouts, spinach, fenugreek sprouts, and apple cider, as well as consumption or

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recreational use of water, direct contact with cattle or animal excreta, attendance at

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agricultural fairs, and recreational use of pastures (King et al., 2012; Rangel et al., 2005).

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Secondary person-to-person transmission, such as within families, day care centers, and

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healthcare institutions, has also been reported (Carter et al., 1987; Rangel et al., 2005;

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Rowe et al., 1993; Spika et al., 1986). A remarkable feature of STEC is its low infectious

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dose; approximately 50–100 bacteria are sufficient to cause disease in healthy individuals

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(Tilden et al., 1996).

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Worldwide, STEC infections cause an estimated three-million acute illnesses annually

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(Majowicz et al., 2014). The average cost per STEC O157 case varies greatly according

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to disease severity: for patients not requiring medical care, the cost is approximately $26,

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whereas those who develop and die from HUS may cost up to $6.2 million per case

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(Frenzen et al., 2005).

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Shiga toxins

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The term “Shiga” originated in 1898, when Kiyoshi Shiga first described the infectious

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agent of epidemic bacterial dysentery, Shigella dysenteriae type 1 (Shiga's bacillus)

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(Trofa et al., 1999). Following this, Keusch et al. (1972) determined that a toxin produced

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by this bacterium caused fluid accumulation and enteritis in ligated intestinal segments

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from rabbits. In 1977, Konowalchuk et al. reported the discovery of a novel cytotoxin in

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cell-free culture filtrates of some E. coli strains. This cytotoxin differed from known heat-

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stable and heat-labile enterotoxins from E. coli. The compound had cytotoxic activity

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against Vero cells and produced markedly different effects from those of heat-labile

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enterotoxin.

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Ten years after the discovery of Stx, which is produced by S. dysenteriae, O'Brien et al.

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(1983) reported that certain strains of E. coli produce a toxin that can be neutralized by

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anti-Stx serum. As a result, E. coli strains that produce Shiga-like toxins were named

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Shiga toxin-producing E. coli, as the Stx1 produced by E. coli is essentially identical at

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the genetic and protein levels to the Stx produced by S. dysenteriae 1. The ability of

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STEC strains to cause severe disease in humans is related mainly to their capacity to

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produce Stx toxins.

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Stxs belong to the AB5 family of protein toxins (Figure 1). These toxins are composed of

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an enzymatically active A moiety and a nontoxic B moiety, which is responsible for

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binding to cellular receptors. The StxB moiety is ring-shaped and pentameric, consisting

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of five identical B subunits (7.7 kDa) surrounding a central pore through which the C-

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terminus of the A moiety is anchored (Fraser et al., 1994; Stein et al., 1992). Each B

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subunit harbors three different binding sites (sites 1–3) that interact with a trisaccharide

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moiety on the glycosphingolipid receptor globotriaosylceramide (Gb3) (Jacewicz et al.,

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1986; Lindberg et al., 1987; Ling et al., 1998; Lingwood et al., 1987). In this way, each B

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moiety can potentially interact with up to 15 Gb3 molecules, resulting in high-affinity

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binding between StxB and Gb3 (Bergan et al., 2012). All Stxs, with the exception of one

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Stx2 variant known as Stx2e, bind to Gb3; Stx2e preferentially binds to the glycolipid

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globotetraosylceramide (Gb4) (Matise et al., 2003).

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To exert its enzymatic activity on target cells, the A subunit of Stx (32.2 kDa) must be

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cleaved into an enzymatically active A1 fragment (27.5 kDa) and a small A2 fragment

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(4.5 kDa) (Garred et al., 1995). After cleavage, the A1 fragment remains attached to the

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A2 fragment via a disulfide bond until the toxin is exposed to the reducing environment

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of the endoplasmic reticulum lumen (Garred et al., 1997). Then, the A1 fragment is

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released and translocates into the cytosol, where it exerts its cytotoxic action, leading to

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cellular death and apoptosis.

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Within the ER, the Stx A1 fragment dissociates from the A2 fragment and the B subunits

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following furin-mediated proteolysis and disulfide bond reduction (Garred et al., 1995;

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Sandvig et al., 2010). From the ER, the proteolytically processed A1 enters the host cell

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cytosol and removes one adenine from the adenosine at position 4324 in the 28S

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ribosomal RNA. This removal inhibits the binding of aminoacyl-tRNA to the 60S

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ribosomal subunit and therefore inhibits cellular protein synthesis (Paton and Paton,

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1998).

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The precise mechanism(s) by which the different Stxs activate apoptosis remain to be

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clarified. Stx1 and Stx2 induce apoptosis and activate stress response pathways in

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endothelial cells. After internalization, Stx2 activates various intracellular stress

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pathways, such as the endoplasmic stress response pathway and the ribotoxic stress

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response pathway (Tesh, 2012). This may lead to apoptosis via the activation of various

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cell death signals, such as the downregulation of the antiapoptotic Bcl2 protein

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(McCullough et al., 2001), the activation of the mitogen-activated protein kinases

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(MAPKs) p38α and c-Jun N-terminal kinase (JNK) (Kitamura, 2008; Smith et al., 2003),

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and the induction of caspase 3-dependent apoptosis (Orrenius et al., 2003; Zong et al.,

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2003).

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The Stx family is divided into two immunologically non-cross reactive groups known as

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Stx1 and Stx2 (O’Brien and Holmes, 1987). STEC strains can express only Stx1, only

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Stx2 or both. The AB5 holotoxin structure is conserved among all Stx family members

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(Nataro and Kaper, 1998; Paton and Paton, 1998).

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Although the Stx family members all share the same holotoxin structure and biological

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activity, differences exist among toxin variants. Stx produced by S. dysenteriae and Stx1

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from E. coli have identical B moieties (De Grandis et al., 1987) and differ at only one

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residue in the A moiety (Strockbine et al., 1988). However, Stx2 is immunologically

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distinct from Stx1, and they share only approximately 56% amino acid sequence identity

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(Calderwood et al., 1987; Jackson et al., 1987).

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There are three Stx1 subtypes (Stx1a, Stx1c, and Stx1d) and seven Stx2 subtypes (Stx2a,

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Stx2b, Stx2c, Stx2d, Stx2e, Stx2f, and Stx2g) (Scheutz et al., 2012).

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Epidemiologically, Stx2 is considered the most important Stx subtype, as the probability

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of HUS development following infections with STEC strains producing only Stx2 is

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higher than that following infections with STEC strains synthesizing only Stx1 or both

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Stx1 and Stx2 (Friedrich et al., 2002). As such, Stx2a, Stx2c and Stx2d are the toxin

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variants most commonly associated with severe outcomes in STEC-infected humans

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(Bielaszewska et al., 2006; Persson et al., 2007). Stx2e is associated with high mortality

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rates in infected pigs; this variant causes Swine Edema Diseases and is responsible for

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significant economic losses in the swine industry (Arimitsu et al., 2013).

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Stx-encoding genes are harbored by lambdoid phages and can therefore be found in free

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phage particles independent of E. coli (Schmidt, 2001). Furthermore, it has been reported

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that the high prevalence of Stx-encoding phages in the environment could lead to the

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generation of new STEC pathotypes via transduction (Cornick et al., 2006; Herold et al.,

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2004; Schmidt, 2001). Although horizontal transfer of Stx-encoding genes occurs at a

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relatively low frequency, the recent large and deadly outbreak of E. coli O104:H4 in

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Germany appears to have been caused by such a recombination event, which resulted in

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the creation of a highly pathogenic hybrid strain (Mellmann et al., 2011; Scheutz et al.,

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2011).

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STEC infection treatments and associated clinical complications

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To focus only on the most up-to-date treatment strategies for STEC infections and STEC-

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associated HUS, we limited the current review to reports of the last major STEC

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outbreak, which took place in Germany in 2011.

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Treatment of STEC infections and STEC-induced HUS remains challenging. There are no

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specific therapies for HUS or vaccines available to prevent the condition (Karmali, 2004).

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In humans, HUS is characterized by the widespread formation of thrombotic

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microvascular lesions in the renal glomeruli, gastrointestinal tract, brain, pancreas, and

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lungs (Fong et al., 1982; Richardson et al., 1988; Upadhyaya et al., 1980). Stxs

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specifically attack small blood vessels mainly in the digestive tract, kidneys and lungs.

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The vascular endothelium of the glomeruli in the kidney is a specific target for Stx, and

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destruction of these filtering structures compromises renal function and causes kidney

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failure and the development of HUS, which is often fatal (Karch et al., 2012)

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The 2011 STEC outbreak in Germany illustrated the lack of specific therapy against

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HUS. In this outbreak, over 4000 individuals were infected with a rare hybrid Stx2-

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producing strain, and more than 900 cases of HUS and 54 deaths resulted (Karch et al.,

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2012). One of the most critical limitations toward successfully containing the outbreak

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was the absence of specific anti-Stx2 therapies.

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The patients in the 2011 Germany outbreak received supportive care to maintain fluid and

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electrolytic levels and dialysis for removal of the toxin from the bloodstream. The

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majority of the patients received plasmapheresis, and a subset of patients received

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eculizumab. A few patients with neurological complications who did not respond to

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plasma exchange and eculizumab were treated with IgG immunodepletion via

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immunoadsorption (Greinacher et al., 2011).

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As described above, Stx is composed of two moieties. The A moiety possesses N-

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glycosidase activity, while the pentameric B moiety interacts with Gb3 receptors on the

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host cell membrane. To exert its cytotoxic activity, Stx invades cells through retrograde

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membrane

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(Mukhopadhyay and Linstedt, 2012). Following the retrograde subcellular transport of

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Stx, the A subunit dissociates from the holotoxin, translocates into the cytosol, and targets

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ribosomes, disrupting protein synthesis and resulting in cell death. Although the

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administration of small molecule inhibitors, such as manganese, in non-toxic doses

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(Mukhopadhyay and Linstedt, 2012) and the use of heterocyclic molecules with central

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imine or benzodiazepine moieties (known as Retro-1 and Retro-2) (Stechmann et al.,

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2010) can inhibit retrograde membrane trafficking and protect mice from toxin-induced

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lethality, these reagents are not currently approved for clinical use.

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Using antibiotics to treat STEC infections poses certain risks and can be contraindicated

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in select clinical situations (Tarr et al., 2005; Wong et al., 2012) because they can promote

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the induction of temperate bacteriophages carrying Stx-encoding genes or lead to the

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release of high quantities of toxins (Kimmitt et al., 2000). The use of antibiotics may

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significantly increase the risk that a STEC infection progresses into HUS (Tarr et al.,

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2005). Bacteriophage production is associated with the induction of the SOS response, a

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bacterial response to DNA damage. SOS-inducing antimicrobial agents, particularly

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quinolones, trimethoprim, and furazolidone, have been shown to induce the expression of

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toxin-encoding genes. Furthermore, when used at levels above those required to inhibit

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bacterial replication, these agents may potentiate the transcription of Stx2-encoding genes

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by up to 140-fold (Kimmitt et al., 2000). According to the referenced study, the use of

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SOS-inducing antimicrobials in clinical practice and animal husbandry may contribute to

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the emergence of STEC-associated diseases.

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For EHEC-associated gastroenteritis, supportive care and intravenous fluid replacement

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are the cornerstones of therapy (Page and Liles, 2013). The administration of drugs with

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anti-motility effects is not recommended because these drugs have been associated with

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an increased risk of developing HUS in some studies (Bell et al., 1997; Piercefield et al.,

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2010). Furthermore, non-steroidal anti-inflammatory drugs should also be avoided

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because of a theoretical risk of worsening gastrointestinal bleeding and/or acute kidney

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injury (Page and Liles, 2013). This issue was partially readdressed during the recent E.

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coli O104:H4 outbreak. Conversely, in vitro studies using outbreak strains have produced

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controversial results, and reports regarding the induction of Stx synthesis following the

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administration of ciprofloxacin have been conflicting (Bielaszewska et al., 2012;

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Corogeanu et al., 2012; Rasko et al., 2010).

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Moreover, the administration of azithromycin was not found to induce Stx production in

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vitro, and its use in patients with HUS has been associated with a shorter duration of fecal

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shedding of bacteria (Nitschke et al., 2012). Therefore, at present, the routine use of

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antibiotics during early EHEC infection is not recommended.

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Recently, Bielaszewska et al. (2012) showed that the antibiotics meropenem,

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azithromycin, rifaximin and tigecycline did not induce phage production or increase toxin

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levels; these antibiotics actually decreased toxin production in vitro. However, further

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studies in animal models and careful analyses of clinical outcomes in patients treated with

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these antibiotics are necessary to determine their potential usefulness for treating humans

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infected with EHEC O104:H4 or other STEC strains.

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No clinical benefits have been associated with other treatments, such as therapeutic

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anticoagulation, administration of fresh frozen plasma or glucocorticosteroids, or the use

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of specific Stx binders (Bitzan et al., 2010).

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During the 2011 outbreak in Germany, many patients with HUS were treated with

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eculizumab. This decision was based on a report attributing the sudden recoveries of three

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children with HUS to treatment with this monoclonal antibody (Lapeyraque et al., 2011).

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According to Davis et al. (2013), the data reported in the above-referenced study

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contained insufficient evidence of eculizumab’s efficacy and therefore its use could not

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be recommended. Other analyses of the O104:H4 outbreak have also concluded that

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eculizumab is of no benefit to infected adult patients (Menne et al., 2012).

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Nanobodies

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Nanobodies are antigen-binding, single-variable-domain proteins derived from naturally

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occurring heavy-chain-only antibodies (Hamers-Casterman et al., 1993). As hydrophobic

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interactions with a light chain are not required, nanobodies are highly soluble,

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physicochemically stable and can be produced with high yields in eukaryotic or

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prokaryotic host organisms.

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Nanobodies combine the desirable features of conventional antibodies with many of the

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desirable properties of small molecule drugs. Like conventional antibodies, nanobodies

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possess high specificities and affinities to a wide variety of antigens. Their potential for

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causing side effects is reduced because of their highly selective binding.

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In addition, the unique structures of the antigen-binding sites in nanobodies enables their

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binding to a wide range of protein epitopes. Other advantages include the feasibility to

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combine VHH domains, which may be directed to the same or different targets to

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enhance their effectiveness. Additionally, nanobodies can be relatively easily produced in

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microorganisms and offer economic advantages compared to conventional mAbs.

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Furthermore, they can be engineered to achieve high thermodynamic, chemical and

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storage stability, as well as high solubility and resistance to proteases (Kolkman and Law,

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2010). These favorable properties have led to the development of several nanobodies for

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use in a wide variety of therapeutic applications (Hassanzadeh-Ghassabeh, 2013; Van

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Bockstaele et al., 2009).

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The single-domain nature and small size of nanobodies enables them to be genetically

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engineered via fusion into multimeric constructs with multiple specificities (Harmsen and

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De Haard, 2007; Muyldermans, 2013). This process, also called “formatting”, may

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involve multimerization of nanobodies via their binding to the same or different targets.

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Nanobody multimers are generated by genetically fusing monomers with short peptide

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linkers. In a virus neutralization assay, Kolkman and Law (2010) observed that the

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neutralization IC50 values for monomeric nanobodies improved by over 4000-fold when

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trimers of the same nanobody were assayed. This avidity effect can also be improved by

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linking two different nanobodies that bind to two different epitopes to create multispecific

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nanobodies, as demonstrated by Conrath et al. (2001). For therapeutic applications, this

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bispecific format is attractive; for example, it can be used to block more than one

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functional epitope on a target antigen and can also extend the in vivo half-life of a

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nanobody.

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Incorporation of a serum albumin-binding monomer into a nanobody may prolong its

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serum half-life from <1 hour up to that of serum albumin, which in humans is

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approximately two to three weeks (Kolman and Law, 2010). Klooster et al. (2007) also

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observed that clearance rates could be drastically reduced by coupling nanobodies to anti-

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human serum albumin. In this way, their usefulness as potential drug candidates could be

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enhanced. The ability to tailor the serum half-lives of nanobodies is critical to their broad

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therapeutic applicability because they are normally rapidly cleared from the human body.

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Conversely, the exceptional stability of nanobodies leads to possibilities for new

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therapeutic applications and alternative routes of administration. Van der Vaart et al.

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(2006) developed nanobodies against rotavirus that could retain functional activity in the

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gastrointestinal tract and were resistant to the acidic environment of the stomach.

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Moreover, these nanobodies reduced the morbidity of rotavirus-induced diarrhea when

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tested in a mouse model.

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Peptides

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Peptides are attractive platforms for the development of therapeutics because they

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combine useful properties, such as high binding affinity, excellent target specificity, low

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toxicity and relatively small mass. However, peptides have short in vivo half-lives, which

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has hampered the development of peptide-based drugs. Thus, the fast elimination of

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peptides from circulation is mainly associated with enzymatic degradation and/or fast

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renal clearance (Pollaro and Heinis, 2010; Werle and Bernkop-Schnürch., 2006).

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Generally, peptides are cleared from the bloodstream within minutes after administration

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(McGregor, 2008; Vlieghe et al., 2010). Peptide elimination primarily occurs via the

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kidney, where glomerular ultrafiltrate is pressed out of the plasma. Glomeruli have an

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approximately 8-nm pore size, and peptides and other molecules with masses below 5

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kDa are completely filtered out (Pollaro and Heinis, 2010).

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To prevent enzymatic peptide degradation, a range of effective, widely applicable

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strategies based on chemical modification of peptides have been developed. Such

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strategies include backbone modification, side chain substitution, D-amino acid

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utilization, cyclisation, and termini modification, among others (Adessi and Soto, 2002;

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Werle and Bernkop-Schnürch, 2006).

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Because proteins larger than 50 to 70 kDa are not rapidly filtered by the kidney, peptides

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and small proteins have been conjugated to long hydrophilic synthetic or natural

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polymers, recombinant polymer mimetics or carbohydrates to increase their

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hydrodynamic volumes (Kontermann, 2009). The most widely employed polymer for

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increasing protein size is polyethyleneglycol (PEG), a molecule composed of repeating

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ethylene oxide units (Jevsevar et al., 2010; Veronese and Pasut, 2005). In addition to

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reducing renal clearance, PEG also stabilizes peptides to protect them from proteolytic

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degradation, increase their solubility and decrease their immunogenicity (Pollaro and

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Heinis, 2010).

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As an alternative to conjugation with synthetic PEG polymers, proteins and peptides can

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be linked to natural polymers, such as polysialic acid (PSA) or hydroxyethyl starch (HES)

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(Gregoriadis et al., 2005).

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To extend their half-lives, peptides have been coupled to albumin and immunoglobulin

338

fragments because the large sizes of albumin and immunoglobulin (67 kDa and 150 kDa,

339

respectively) prevent fast renal clearance (Pollaro and Heinis, 2010; Sato et al., 2006).

340

The above-described studies indicate that additional strategies aimed at increasing the

341

half-lives of peptides will improve the therapeutic efficiencies of both existing and novel

342

peptide drugs.

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Phage display and Stx neutralizers

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First reported by Smith in 1985, phage display technology has been key to the

346

development of antigen-specific peptides and proteins. Phage display is an in vitro

347

selection technique that enables peptides or proteins with highly specific properties to be

348

enriched from a large collection of variants. Used for the recognition of specific target

349

molecules and biomarkers, phage display has yielded economic, rapid, and efficient

350

applications in fields such as vaccine development, enzyme inhibition, inflammation

351

reduction, and cancer research (Lee et al., 2013). Moreover, phage display has become

352

one of the most powerful drug discovery platforms and can also be used to engineer many

353

of the attributes associated with successful drugs. Such attributes include potency,

354

specificity, cross-reactivity and stability (Nixon et al., 2014).

355

The replication scheme and structure of Ff filamentous phage from E. coli have been

356

extensively used in phage display technology, and a wide range of nanotechnology

357

applications have been derived as a result. These filamentous phages are the most

358

productive phages in nature. The best studied of such phages include the F pilus-specific

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or Ff phages, such as f1, M13 and fd (Rakonjac et al., 2011).

360

Phage display is a powerful tool for the selection of peptides and protein domains,

361

including antibodies. One advantage of this methodology is the direct physical link that

362

exists between phage phenotype and genotype. Linking a polypeptide or protein on a

363

phage surface (phenotype) to its encoding DNA (genotype, which is integrated into the

364

phage genome) allows researchers to screen large recombinant peptide and protein

365

libraries for the presence of highly specific clones using the discriminative power of

366

affinity purification (Konthur and Walter, 2002; Tikunova and Morozova, 2009).

367

The goal of phage display library screenings is the identification, selection and isolation

368

of “rare” variants with specific biological features. Initially, engineered phages must be

369

assembled and released from host cells in sufficient numbers to ensure that a desired

370

variant is present within and can be selected from the library. Generally, success in

371

screening depends on the affinity and specificity of a peptide for its ligand, as well as the

372

number of copies of the peptide per phage particle (Rakonjac et al., 2011).

373

The basic method of using affinity screening to select isolates from phage display

374

libraries is often referred to as “biopanning” (Parmley and Smith, 1988). During

375

biopanning, a target ligand is immobilized on a solid support and then exposed to a phage

376

display library to facilitate the binding of specific variants. Multiple rounds of washing

377

are performed to eliminate adherent but non-binding phages, and the remaining ligand-

378

bound variants are then eluted. To overcome non-specific binding of phages to the matrix

379

surrounding the ligand and to enrich the number of binding variants, at least three rounds

380

of biopanning are recommended to eliminate the “background” associated with non-

381

specific binders (Rakonjac et al., 2011).

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The first phage display libraries of peptides and antibodies were reported in the early

383

1990s (Barbas et al., 1991; Clackson et al., 1991; Cwirla et al., 1990; Devlin et al., 1990;

384

Marks et al., 1991; Scott and Smith, 1990). Following these reports, the number of phage

385

display applications for protein and antibody engineering quickly increased.

386

Two types of phage display libraries have been used for the majority of reported

387

applications: random peptide libraries and antibody libraries.

388

Random peptide libraries are typically used to identify peptide candidates that bind to a

389

target of interest. In a pioneering work, Scott and Smith (1990) constructed short, random

390

peptide libraries and screened them for the ability to bind to a ligand of interest. Peptide

391

libraries contain peptides of variable lengths and conformations; these peptides are

392

typically displayed in a loop-constrained conformation induced by two flanking cysteine

393

residues or in linear or bicyclic conformations (Chen and Heinis, 2015; Felici et al.,

394

1993).

395

In the early 1990s, antibody phage display led to the use of in vitro phage affinity

396

selection as an interesting alternative to conventional immunization and hybridoma

397

production for the successful isolation of monoclonal recombinant antibodies that

398

recognize an antigen of interest. These early reports showed that the variable domains of

399

the heavy (VH) and light (VL and VK) chains of antibodies can correctly fold and

400

recognize their cognate antigens when expressed in E. coli and fused to pIII or pVIII

401

proteins from Ff bacteriophages (Barbas et al., 1991; Kang et al., 1991; McCafferty et al.,

402

1990).

403

Using phage display to identify agents that neutralize bacterial toxins has significantly

404

increased in popularity in recent years. Overall, the majority of scientific research

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centered on phage display can be allocated into random peptide phage display studies and

406

antibody phage display studies (mainly consisting of libraries of filamentous phages

407

displaying Fab antibody fragments or nanobodies).

408

Herein, we provide a brief overview of the progress achieved through the use of phage

409

display technology for the development of potential therapeutic agents to treat the clinical

410

manifestations associated with Stx-induced cytotoxicity.

411

One of the first reports regarding the use of phage display to combat Stxs was published

412

by Han et al. in 1999. In this report, receptor antagonists were selected using phage

413

libraries, and phage clones capable of binding to the StxB subunit were isolated. These

414

authors also described the selection of a 15-mer peptide (A12) displayed on fd

415

bacteriophages that efficiently competed with the Gb3 receptor for binding to StxB.

416

Additionally, the authors reported that peptide A12 inhibited the cytotoxicity and

417

enterotoxicity of Stx from Shigella dysenteriae type 1.

418

Later, Miura et al. (2004) used self-assembled monolayers (SAM) of Gb3 trisaccharide-

419

mimics to select candidates from dodecapeptide-expressing phage display libraries.

420

Previously, Oldenburg et al. (1992) had already demonstrated the usefulness of random

421

peptide libraries for the selection of peptide ligands for sugar-binding proteins. Thus,

422

after three rounds of biopanning and analysis of the isolated phage sequences, Miura et al.

423

selected two peptides containing a FHENWPS consensus fragment and observed that

424

these peptides inhibited Stx1 binding to SAM.

425

In the same year, Inoue et al. (2004) reported cloning Stx1-neutralizing monoclonal

426

antibodies, named 5-5B, from hybridoma cells using phage display techniques. The

427

recombinant Fab fragments were characterized and showed binding activity against Stx1,

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but not against Stx2. Furthermore, the 5-5B Fab fragments exhibited neutralizing

429

activities against Stx1-induced cytotoxicity

430

Zhang et al. (2004) developed pentamerized single-domain antibodies (sdAb) isolated

431

from a naive llama sdAb library and linked them to an oligomerization domain to

432

generate high-avidity antigen-binding reagents. In this report, an sdAb was fused to the B

433

subunit of Stx1; this complex self-assembled to form an sdAb-homopentamer with high

434

avidity. Although the focus of this report was not the identification of Stx-neutralizing

435

antibodies, the authors described dramatic increases in affinity for the Stx1B-immobilized

436

antigen. In addition, it was reported that these pentabodies were expressed in high yield in

437

E. coli and exhibited enhanced thermostability and resistance to proteases.

438

Two years following the above-referenced report, Bao et al. (2006) used recombinant

439

StxB to screen dodecapeptide phage libraries for candidates capable of binding to this

440

subunit. After four rounds of biopanning, three phages clones, named A3, A6 and A9,

441

that could bind to the target were isolated. Subsequently, when Stx was administered with

442

the A6 phage clone and inoculated in BALB/c mice, survival of the mice increased by

443

33.3%.

444

Based on results reported by Miura et al. (2006), Yamada et al. (2006) designed

445

bifunctional short peptides by fusing nuclear localization signal peptides with a

446

globotriaoside (Gb3)-mimic peptide that was developed using a phage display approach

447

(Miura et al., 2006). The authors noted that one engineered peptide that bound strongly to

448

Stx1 and Stx2, called Fusion peptide 2, expressed antibacterial activity against E. coli and

449

Staphylococcus aureus and neutralizing activity against Stx1 cytotoxicity in HeLa cell

450

cultures.

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In 2007, Stone et al. (2007) developed a potential neutralizer for Stx cytotoxicity using

452

pentamerized sdAbs (Zhang et al., 2004) isolated from a naive llama phage display

453

library. They reported that two pentameric sdAb neutralizers, VTI-1 and VTI-3, bound to

454

the wild-type Stx1B subunit with high affinity and respectively inhibited the cytotoxic

455

activity of Stx1 by approximately 90% and 40% when evaluated using Vero cell-based

456

assays. However, these pentabodies could not prevent lethality in mice challenged with

457

lethal doses of Stx1; as the concentration of pentabodies required for in vitro

458

neutralization of this toxin was extremely high, it is likely that an effective dose could not

459

be reached in vivo.

460

In 2011, Neri et al. isolated single-chain variable fragments (scFv) against Stxs by

461

screening a naïve phage display antibody library. They described one antibody, B22, that

462

neutralized Stx1-mediated cytotoxicity in HeLa 229 cells. However, no neutralizing

463

activity was observed when varying doses of Stx2 were assayed. Additionally, only the

464

binding of His-tagged Stx1B to its cellular surface receptor was completely inhibited by

465

the B22pp antibody.

466

Following the above, Tremblay et al. (2013) reported the use of a VHH (single-domain

467

fragments or nanobodies) phage display library to identify VHHs capable of binding Stx1

468

and/or Stx2. They observed that the majority of isolated Stx-binding VHHs recognized

469

the B subunit and neutralized the target in Vero cell assays. Additionally, the authors

470

described the identification of one VHH that recognized both Stx1 and Stx2. After

471

administering a single VHH-based neutralizing agent consisting of linked VHH

472

heteromultimers (VNA) together with an anti-tag monoclonal antibody engineered in

473

VNAs (effector Ab or efAb), they observed toxin clearance and protection against

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lethality in mice given lethal doses of Stx1 and Stx2.

475

Recently, Lo et al. (2014) reported the use of nanobodies expressed on llama VHH-phage

476

display libraries to select candidates capable of neutralizing the Stx2e variant. The

477

referenced study showed that the NbStx2e1 nanobody conferred strong neutralizing

478

activity against Stx2e cytotoxicity in Vero cell-based assays. Structural analysis revealed

479

that this nanobody competed for the glycolipid receptor-binding site.

480

During the same year, Bernedo-Navarro et al. (2014) described the use of peptide phage

481

libraries displaying linear (12-mer) and loop-constrained (7-mer) oligopeptides to isolate

482

peptides capable of neutralizing Stx1 and Stx2 (Bernedo-Navarro et al., 2014). In this

483

study, the Gb3 receptor was used as a target for biopanning, and three Gb3-binding

484

peptides, called PC7-12, P12-26 and PC7-30, were identified. These peptides inhibited

485

cytotoxicity in Vero cell-based assays and competed efficiently by binding to Gb3

486

receptors; however, only peptide PC7-30 inhibited Stx1-induced lethality, but not Stx2-

487

induced lethality, in mice.

488

More recently, Luz et al. (2015) developed recombinant antibody Fab fragments targeting

489

Stx2 that were selected from human synthetic antibody libraries displayed on M13

490

bacteriophages. The Fab fragment described by these researchers could inhibit Stx2

491

toxicity by approximately 70% in vitro.

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Conclusions

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Currently available information suggests that phage display technology will remain a

495

powerful tool for the development of therapeutic antibodies and drug-like molecules with

496

enhanced biological features. With additional future improvements, this versatile and 22

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flexible tool offers an excellent approach not only for the production of engineered

498

peptides and antibodies against Stx biological activity but also for the development of

499

new therapeutic agents against other toxins. Another interesting aspect of phage display

500

technology is its cost effectiveness in the in vitro production of antibody fragments, such

501

as binding fragments (Fab) or single-chain variable fragments (scFv).

502

Collectively, the above-described peptides and nanobodies obtained through phage

503

display technology exhibited different levels of Stx-neutralizing activity. Although these

504

molecules could inhibit Stx-mediated cytotoxicity both in vitro and in vivo, some of the

505

molecules should be structurally optimized and modified in other ways to extend their

506

half-lives and enhance their biological effects. Developing efficient therapeutic

507

candidates against Stx-induced damage is an extremely challenging task, as demonstrated

508

by the more than three decades of research that has gone into this field. The complexity

509

of the interactions between Stxs and their cellular receptors in combination with the quick

510

development of severe clinical manifestations has hampered the successful identification

511

of efficient neutralizers. However, despite the lack of a specific and well-established

512

therapy for these toxins, phage display technology has proven to be a powerful tool for

513

improving the identification of these urgently needed reagents.

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Acknowledgements

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We thank Miguel Montoute for reviewing this document.

517

The corresponding author is a Post Doctoral Fellow of Fundação de Amparo à Pesquisa

518

do Estado de São Paulo (FAPESP).

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Figures

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Figure 1. Shiga toxin structures (based on Fraser et al., 1994).

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a. Shiga toxin (Stx) organization.

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b. S. dysenteriae Stx structure showing the A and B moieties (PDB ID: 1DM0).

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c. Ring-shaped StxB structure consisting of five identical B monomers (PDB ID:

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1DM0).

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Figure 2. General workflow for phage display screening of diverse targets.

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a. Specific phage libraries are chosen and can be customized by genetic engineering.

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b. Phage display permits the screening of a wide variety of targets against phage

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libraries.

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c. After selecting target-binding phage clones, eluted phages are amplified in a proper host and put through binding/amplification cycles to enrich the pool in favor of binding clones.

d. After 3-6 biopanning cycles, individual clones are characterized by DNA sequencing to identify binding consensus sequences. 46

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e. Peptides/antibody fragments are synthesized after characterization and in silico sequence analysis. f. Peptides/antibody fragments are biologically characterized in vitro and in vivo, and subsequent structural analysis can be performed.

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We would like to inform that this manuscript is a review of the scientific literature

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published by different groups of researchers; therefore animals were not used.