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11.2.2.1 Nuclear Magnetic Resonance NMR has become a favored technique
for screening fragment libraries following the pioneering work of SAR by NMR at
Abbott Laboratories [15b, 33]. Not only is it capable of detecting weak interactions
with high reliability, it may also provide information about the site of interaction,
assisting in the ranking and exploitation of the fragment hits. Furthermore, it monitors
binding of both partners directly in free solution, without interference from immobilization or detection artifacts, by following the change of intrinsic NMR spectral
parameters that are highly sensitive and selective to the binding event between ligand
and receptor. Ligand binding can be monitored by observing the effects on the NMR
spectra either of the target [34] or of the ligand itself [35].
Target-based NMR screening relies on the detection of perturbations of target
resonances on ligand binding [36]. The most well-known technique is the SAR by
NMR method. Ligands are identified by monitoring alterations of target signals in
a2D
1H–15
N correlation spectrum. This method requires large quantities of isoto-
pically (
15
N)-labeled protein (50–200 mg, with protein solubility between 0.1 and
1 mM) [37]. This method has the benefit (provided that sequence-specific resonance
assignments have been obtained) of high reliability, being able to identify the binding
site on the target, offering K
d
information, readily distinguishing specific from
nonspecific interactions, and assessing whether any significant conformational
changes occur upon binding. The incorporation of
13
C into the target protein may
also be used for the detection of binding of small molecu les to targets [38].
Another method for screening larger protein targets is site-selective screening
with labeled amino acid pairs [39], which relies on the sequence-specific labeling
technique [40]. Using this labeling strategy, it is possible to selectively screen the
ligand without sequence-specific assignment s, and the chemical shift perturbations
upon binding of a potential ligand are easily detected.
The limitation of target-based NMR screening is that the size of targets that can be
observed to molecular weights is usually <100 kDa even if techniques like transverse
relaxation optimized spectroscopy (TROSY) [41] or cross-relaxation induced
polarization transfer (CRIPT) [42] are applied.
Ligand-based NMR screening relies on changes of the ligand signals when binding
to the target [35, 43]. Several methods have been developed. One is the detection of an
altered hydrodynamic property (i.e., molecular tumbling rate or diffusion rate) upon
target binding [44]. When a small ligand binds to a macromolecule, its apparent rates
of diffusion and reorientation are decreased. Such decreased diffusion rates can be
measured by gradient-enhanced NMR spectroscopy. The decreased rate of molecular
tumbling (increased rotational correlation time) is dir ectly manifested in an increased
transverse relaxation rate of the NMR signals, which can also be easily measured.
The transferred nuclear overhauser enhancement (NOE) technique also falls into this
class [45]. It relies on detecting intraligand NOEs that develop in the bound state,
where the dipole–dipole interaction caused by the decr eased molecular tumbling rate
is much stronger than in a free state of a free ligand.
A second ligand-based NMR screening method involves detection of the transfer
of an NMR signal between target and ligand. Methods that represent this class
of experiments are NOE pumping [46], saturation transfer difference (STD) [47], and
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water–ligand observed via gradient spectroscopy (waterLOGSY) [48]. These techniques are closely related in that they all rely on dipole–dipole interactions between
ligand–target spins. NOE pumping and waterLOGSY represent coherent methods
because net magnetization is transferred. In this context, STD is considered an
incoherent method because it relies on the transfer of saturation. This set of
experiments represents the most attractive NMR-based techniques for binding
detection currently available because it enables a lower protein concentration than
other techniques.
A third ligand-based method is NMR fragment screening detecting interligand
NOEs [49]. This method solely detects the signals between two small-molecule
fragments. When two fragments bind to the target at neighboring sites, such that their
hydrogen atoms are less than 5–6 A˚apart, a transferred NOE can be detected between
the hydrogen atoms by 2D NMR spectroscopy. Another ligand-based method
involves the use of paramagnetic spin labels, such as 2,2,6,6-tetramethyl piperidine1-oxyl, to increase the relaxation of nearby spins [50]. The spin label is covalently
attached to either a ligand with a known binding site or a reactive amino acid side
chain at the edge of the desired binding pocket. When another fragment binds to
the target within a distance of less than 15–20 A˚from the spin label, its
1
H signals can
be selectively weakened. The fifth ligand-based method uses
19
F NMR to detect
binding of fluorinated compounds to a target [51]. The wide chemical shift range and
the simple signal pattern for the fluorinated molecules can facilitate the direct
assignment of compounds within a mixture that bind to a protein. However, this
technique suffers from intrinsic low sensitivity. It can be overcome by the use of an
optimized cryogenic
19
F probe and the incorporation of magnetically equivalent
fluorine atoms in ligands through the use of
19
F label of the CF3group [52]. The last
ligand-based method described here is target-immobilized NMR screening, which
immobilizes a target on a solid support and allows rapid characterization of the ligandbinding site [53]. This approach holds the potential for fragment screening of integral
membrane proteins.
Contrast to target-based NMR, the protein concentration in ligand-based NMR
screening is in the low micrometer range; therefore, the protein consumption is
substantially lower. Ligand-based methods do not require protein labeling, so there is
no upper limit on the size of the protein. However, the information content is lower
(there is, e.g., no direct information on the site of binding, although this can sometimes
be obtained indirectly) [54]. Most ligand-detection methods are usually limited to low
and medium affinities because of the need to suppress the signals of the target
molecule. Ligands that bind too tightly are indistinguishable from the target, and thus
are suppressed as well, resulting in a false negative. In contrast, nonspecific binding
can result in the appearance of false positives, so having access to a positive control
(a known competing ligand) is almost a prerequisite to allow for the discrimination
between specific and nonspecific effects [37].
11.2.2.2 X-Ray Crystallography X-ray crystallography can yield the most
complete picture of fragment binding to its target. Similar to NMR, crystallography
has the advantages of defining the ligand-binding sites and the binding orientations of
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the molecular fragment with more certainty. With advances in protein production
and crystallization, and the degree of automation associated with crystal manipulation, X-ray data collection, and data processing [55], solving crystal structures is
increasingly high throughput in recent years [16]. The use of synchrotron radiation
sources to generate high-intensity X-rays has enabled many novel structures to be
determined at unprecedented rates [56]. The improvement of these techniques has
allowed X-ray crystallography to become a primary screening tool for fragment
libraries.
Ringe and coworkers alleviated an initial concern that small weakly binding
fragments might have insufficient affinity to yield well-solved electron density in
a crystallographic structure by demonstrating that even simple organic solvent
molecules could bind to specific sites on protein surfaces [57]. Stroud and coworkers
demonstrated that individual fragments of the substrate dUMP bound to the enzyme
thymidylate synthase in a position similar to that of the full substrate [58]. A survey of
the Protein Data Bank has concluded that two structurally similar ligands belonging to
the same series in a drug design project can safely be assumed to occupy the same 3D
position in the binding site [59].
One of the first research groups to address fragment-based crystallography
screening was Verlinde and coworkers. They described a successful fragment-based
design of an inhibitor for triose phosphate isomerase (TIM) from Trypanosoma
brucei [60]. Nienaber and coworkers described a CrystaLEADS (crystallographic
screening for lead compound fragments) approach, which uses a library of 10,000
compounds divided into cocktails of 100 compounds [61]. The cocktails were
assembled such that each obtained compounds of diverse shapes. SAR by X-ray
was proposed by a group at Aventis [62]. A group at Astex developed a fragmentbased crystallographic screening procedure, Pyramid , which has been widely used
in the generation of their lead structures [63].
Crystallography-based screening requires 10–50 mg of target protein with a purity
of >95% [64]. Efficient fragment screening requires the soaking of a mixture of
fragments (a cocktail) into the preexisting crystals of the target protein at high
concentration (>10 mM) in the presence of organic solvents (usually DMSO) [65].
The fragments in each cocktail are usually selected to be both highly soluble and also
shape diverse to assist interpretation of electron density. The crystals typically need
to be able to tolerate cosolvents and high-solute concentrations. After collection of
the X-ray data, the identification of the active fragments from the cocktail can be
automated and accelerated by software tools such as Quanta from Accelrys [66] and
AutoSolve from Astex [67].
Despite many challenges, a crystallographic fragment screen can be very powerful
because it directly furnishes the structural information required for the medicinal
chemistry elaboration of the identified fragment hits. Today many labs have made
crystallographic screening a core component of the fragment-based lead generation
platforms [62, 68]. The major drawback to this method is that it can only be applied
to targets that form crystals suitable for cocrystallization or soaking experiments, and
this level of robustness is not easy to achieve for many protein targets. This limitation
also means that important membr ane-bound targets (such as GPCRs) require different
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methodologies. High concentrations of ligands sometimes used in soaking can give
false positives, as most compounds (including very weak binders) might bind to target
proteins.
11.2.2.3 Mass Spectrometry The speed and high sensitivity of mass spectrometry
(MS) make it an attractive approach to protein–ligand binding studies. Two main
approaches have been applied to the discovery of weak binding ligands. A method,
called SAR by MS, was developed to detect the binding of noncovalent weak binding
fragments to DNA or RNA by electrospray ionization mass spectrometry (ESIMS) [69]. Complexes with nucleotides are rich in H-bond interactions, making them
stable in the gas phase and ideal for study by MS. By optimizing the ionization and
desolvation processes, the researchers were able to characterize low-affinity complexes (in the millimolar range) formed between RNA/DNA and small molecules. On
the basis of the observed mass and abundance of the complexes, the researchers were
further able to directly determine both binding affinity and stoichiometry.
Another technique utilizing MS, called tethering, has also been used to identify lowmolecular weight fragments that interact with a protein target at a specific site [70]. The
technique relies on the formation of a disu lfide bond (e.g., a tether) between the
fragment and a cysteine residue in the targetprotein. Ifa nativecysteine does not exist in
the region of interest, one can be inserted by site-directed mutagenesis. The target
protein is exposed to a library of disulfide-containing fragments and fragments with the
greatest affinity for protein sites in the vicinity of the cysteine fo rm the most stable
disulfide bonds. Disulfides are rapidly detected and identified by mass spectrometry,
such as ESI-MS which is sufficiently gentle forasulfur–sulfurbondtoremainintact
during ionization. Performing the screening experiments under partially reducing
conditions ensures that the intrinsic binding properties between fragment and target,
rather than thiol reactivity, drive the selection process. However, this method
requires aprioriknowledge of the binding site, because the cysteine has to be
located in or close to the active site. The demand to generate cysteine-containing
mutant proteins may have potential limitations for the application.
11.2.2.4 Surface Plasmon Resonance Surface plasmon resonance (SPR) is an
optical technique that uses the evanescent wave phenomenon to measure changes in
the refractive index in the immediate vicinity of the surface layer of a sensor chip [71].
SPR is observed as a sharp shadow in the reflected light from the surface at an angle
that is dependent on the mass of the material at the surface. The SPR angle shifts
when molecules bind to the surface and change the mass of the surface layer. This
change in resonant angle can be monitored noninvasively as a plot of resonance signal
(proportional to the mass change) versus time, which allows the study of the
interaction of proteins with ligands in real time.
A typical SPR-binding experiment consists of two phases: the binding of the
soluble analyte to the sensor and the dissociation of the analyte on rinsing with
analyte-free solution. By fitting kinetics from the association and dissociation phase
to appropriate binding models, the corresponding kinetic rate constants k
on
and k
off
(and, hence, Kd, which equals k
off/kon
) may be calculated. Another approach is the
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direct determination of Kdby analyzing equilibrium-binding data generated at
different analyte concentrations.
Because of the high sensitivity of modern SPR sensors, SPR has been shown to be
suitablefor the detectionof low-molecular weight analytesand low-affinity interactions,
which is a necessary condition for screening of fragment–protein interactions [72].
One often-discussed issue of SPR-based biosensors is the necessary immobilization of one of the binding partners and the tethering of molecules to surfaces may
affect the binding constants measured. However, it has been evident that equilibrium,
thermodynamic, and kinetic data from surface experiments can mirror those obtained
in solution, if SPR biosensor experiments are carefully designed [73]. Numerous
immobilization schemes for the formation of SPR sensor surfaces and their applications have been designed [74]. For example, the use of mediating sensor layers,
such as streptavidin layers on gold and the subsequent attachment of biotinylated
molecules, such as antibodies or DNA, have been successfully applied. Monoclonal
antibodies can be covalently attached to the solid support. A fusion protein containing
an antigen tag and a target protein can couple to the surface through the antibody–
antigen interaction. The direct immobilization of DNA sequences on gold via the thiol
moieties has been demonstrated.
Commercially available SPR sensors for coupling of target proteins include
protein A surfaces, carboxymethylated dextran surfaces for covalently coupling to
a variety of analytes exhibiting active groups, lipid bilayers [75] for the immobilization of membrane proteins, and nitrilotriacetic acid-coupled surfaces to immobilize
His-tagged proteins.
Low consumption of the target protein and the convenient accessibility to
experiments are the advantages of fragment- based SPR screening, coupling the target
protein onto the solid support. However, the potential drawback of this methodology
is that the background signal from unspecific binding might be strong because the
binding of low-molecular weight fragments to protein does not change the SPR angle
shifts much.
Both the target protein and the ligands (fragments) have been immobilized on
the sensor surface [18]. One of the most successful immobilization strategies for
biomolecules on sensor surfaces is the formation of self-assembled monolayers on
gold or glass surfaces and subsequent covalent attachment of ligands (or fragment)
of interest using appropriate linker chemistry. The advantage of this methodology is
that the highly ordered monolayers allow precise control over the density of
immobilized ligands and can be designed to minimize unspecific protein adsorption.
However, the orientations of ligands have been predefined during coupling to the solid
support, which might not be able to produce the optimal binding event with the target
protein. Furthermore, this method limits to the opportunity to design a fragment
library for a specific target because all ligands have to be coupled to the solid support.
11.2.2.5 Biochemical Assays
High Concentration Screening Inherited from HTS, high concentration screening
(HCS) is a HTS application to the identification of fragment hits with low-molecular
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weights. Due to the low binding affinity to its target, fragments are generally required
to be used at high compound concentrations, typically 0.1–2 mM, which is close to
the aqueous solubility limit for a substantial number of compounds. The value of
biochemical assays for fragment screening has been initially questioned based on
experience with the traditional HTS, where compound concentrations of 10 mM can
be associated with false positive (or negative) hits. Four typical sources of false
positive (or negative) hits in the HTS functional assays are (1) readout distor tion
by colored, autofluorescent, or quenching compounds [76], (2) readout distortion by
a limited compound solubility in aqueous solution with resulting precipitation [77],
(3) target inhibition due to nonspecific aggregation [78], and (4) irreversible inactivation of the protein target [79].
In spite of these problems and challenges, HCS is attractive to many fragmentbased ligand discovery campaigns because the HTS facilities have been established,
and HCS is fast, convenient, and its outcome is actually the endpoint of the biological
assays (such as K
i
or IC50). HCS can be cost- and resource-effective, provided that the
assay platform remains robust and trustworthy. An increasing number of attempts
using bioassays to screen fragment collections have been reported. The standard assay
and screening techniques such as ELISA [80], absorbance [81], and radiometry
[68c, 82] have been reported successfully to screen fragment collections. Among
them, ELISA-based fragment screening is of particular interest because it may be
advantageous, as the excessive compound concentration is significantly reduced by
the washing steps before detection. However, these studies have only been conducted
on very small collections of fragments, and the selection of fragment hits is prima rily
based on the medicinal chemistry-driven hypothesis.
High-sensitivity microscopic fluorescence techniques such as confocal fluorescence correlation spectroscopy have been adopted for fragment-based HCS, which
can allow hit thresholds lowering to 10–20% inhibition [83]. This technique has
successfully identified inhibitors of Hsp90 [84].
There is no inherent approach to judge the authenticity of a target-specific active
compound because there are a number of pitfalls that need to be considered when
running a HCS campaign. False positives must be removed as quickly as possible to
minimize the risk of carrying them forward. The methods to support their removal
include (1) ensuring that the screening library is of a high standard (removal of
promiscuous hits and ensuring high solubility), (2) using additional well-designed
assays to weed out false hits and confirming the functional biological data, (3)
subsequently using structural biology or biophysics-based screenings to determine
the binding constants, (4) using mutant information if reachable, and (5) further
analog screening to confirm SARs.
Substrate Activity Screening Ellman and coworkers developed a substrate-based
fragment identification method, called substrate activity screening (SAS) [85]. This
method addresses two key challenges in fragment-based screening (1) the accurate
and efficient identification of weak binding fragments and (2) the rapid optimization
of the initial weak binding fragments into high-affinity compounds. A typical SAS
has three steps (1) a library of substrates consisting of the substrate–catalytic
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functionalities and diverse, low-molecular weight fragments are screened using
a single-step, high-throughput fluorescence-based assay; (2) the activity of the
substrate is rapidly optimized by rapid analog synthesis and evaluation; and (3) the
optimized substrates are converted to inhibitors by direct replacement of the
substrate–catalytic functionalities with inhibitor pharmacophores, which match
the catalytic residues in the active site.
In SAS, both an active enzyme and productive active-site binding are required
for the catalytic function. The electronic and steric effects of the substituents in the
vicinity of the catalytic site have to be compatible with the catalytic function of
the target enzyme so that the enzyme is able to identify the substrates. However, SAS
has some prominent advantages such as (1) being able to detect weak binding
fragments, (2) being high throughput and straightforward to perform, and (3) the
catalytic substrate turnover results in signal amplification, and therefore even very
weak substrates can be identified at concentrations where only minimal binding to the
enzyme occurs.
11.2.3 Approaches from Fragment to Lead Structures
In HTS screening, the transformation of a hit into an attractive lead compound is often
achieved by adding the more likely potency-retaining (or increasing) hydrophobic
substituents [9, 86]. This leads to increases in molecular weight and often also
lipophilicity, which frequently result in negative effects on molecular properties, such
as solubility or metabolic stability. Fragment hits derived from fragment-based
screening are of high quality in terms of physicochemical properties; therefore, these
hits often have superior molecular and ADME/Tox properties compared with hits from
HTS. After the fragment hits are identified, the next step is to convert fragment(s) into
a lead structure and maintain the drug-like properties of the generated molecule. There
are three general strategies for converting fragments into a drug-like lead structure:
fragment evolution, fragment linking, and in situ fragment assembly (Figure 11.1).
11.2.3.1 Fragment Evolution Fragment evolution (Figure 11.1a) is analogous to
the standard hit-to-lead medicinal chemistry optimization process, requiring addition
of other functionalities to a preexisting template to improve the interactions with the
target. The basic requirement for fragment evolution is that the preexisting template
should be a small molecule (called “anchor”) that has plenty of diverse opportunities
for increasing potency before exceeding the boundaries defined by Lipinski’s “rules of
five.”
After the initial fragment hit is identified, there are two common strategies used for
fragment evolution (1) if the target three-dimensional structure is accessible, X-ray
crystallography or NMR spectroscopy can be used to determine the binding information of the initial hit to its target. Structure-base d ligand design approaches are used
to accelerate the evolution progress [87]; and (2) when the structural information
of the target is unavailable, the screening of appropriate analogs of the original hit
would be performed to establish a structure–activity relationship (SAR) to guide the
medicinal chemistry efforts.
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Fragment evolution has proved to be the most popular and effective approach from
fragment to lead. One particular application of fragment evolution is to enhance
properties other than the inherent potency of the original molecule or to deal with
some specific issue, for example, an ADME liability.
11.2.3.2 Fragment Linking Fragment linking (Figure 11.1) involves joining two
fragments that have been identified to bind at adjacent sites in a given target protein.
The potency increase achievable from optimally linking two fragments can be an
approximate of an additive effect of two fragments (i.e., the free energy of binding of
the joined molecule is approximately equal to the sum of the free binding energies
of the binding of two fragments) [58, 88], or in some cases, it can even exceed the sum
of the component fragments (Figure 11.2) [89]. In these cases, although the linkers do
not bind to the target protein directly, they assist the optimal binding of fragment hits
to the target receptor. One extreme case is biotin–avidin binding [90]. In either of
these cases, it requires the negative contribution from the linkers to be minimal and the
loss in rigid-body entropy on binding of all fragment hits to the enzyme to be very
small. An analysis of the experimental energetics associated with optimally linked
fragments has suggested that the rigid-body entropy loss on protein binding constitutes a barrier of around three orders of magnitude to the binding affinity. This
barrier is esse ntially independent of molecular mass that implies that there should be
Figure 11.1 Three general strategies for converting fragments into a lead structure. (a)
Fragment evolution. Fragment 1 binds to the receptor at one site. It is evolved by the addition
of more fragments to create good contacts with the active site surface and then grow into
a second pocket of the active site. (b) Fragment linking. Fragment 1 binds to the receptor at one
site. Fragment 2 binds to the receptor at an adjacent site. The two fragments are joined together
by a linking group that allows the lead molecule to span both sites. (c) In situ fragment
assembly. Fragments 1 and 2 bind to receptor sites simultaneously with two reactive groups
positioned within conformational reach of each other. The lead molecule is formed in the active
site by the chemical reaction between the two reactive groups.
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FRAGMENT-BASED DRUG DESIGN: CONSIDERATIONS FOR GOOD ADME PROPERTIES
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a superadditive effect when two small molecular-mass fragments are linked in an
optimal fashion [91].
However, the linking step can be very difficult to achieve. Th e linker has to be of
just the right length and the right conformation to be able to link fragments so that
they can reach their respective binding sites in an optimal way. Access to structural
information is essential for this approach to succeed since it can avoid the otherwise
necessary and very large combinatorial and random search to find the effective linking
scheme. As a consequence of these complications, there are relatively few examples
of successful fragment linking in the literature that have resulted in lead-like hits with
reasonable compound physicochemical properties and biological activity.
11.2.3.3 In Situ Fragment Assembly In situ fragment assembly (Figure 11.1c)
entails the use of reactive fragments that link together to form an active inhibitor in the
presence of a protein target. This approach can be considered in a wider context of
target-guided synthesis (TGS), an umbrella term that covers a variety of systems that
feature small-molecule synthesis orchestrated by a large biomolecule. The essence of
in situ fragment assembly is that the protein serves as a template for synthesis and
thermodynamically selects the combinations of fragments that can be converted to
a larger and more potent ligand. The biological activities of the larger ligand are then
confirmed using the standard biochemical assays. Currently, there are three in situ
fragment assembly techniques: (1) dynamic combinatorial chemistry (DCC) [92], (2)
tethering with extenders [17a] , and (3) in situ click chemistry [93]. Among them,
dynamic combinatorial chemistry and tethering with extenders are the thermodynamically controlled processes, while in situ click chemistry is a kinetically controlled process.
Dynamic Combinatorial Chemistry DCC was developed from combinatorial
chemistry.The principal difference between DCC and the conventional combina torial
chemistry is that the reaction linking the building blocks together in DCC is reversible
causing an ongoing interchange between the different members of the dynamic
combinatorial library under thermodynamic control. A dynamic combinatorial
library is able to respond to a molecular recognition event owing to the presence
of the target protein, which can stabilize a particular member of the library and induce
a shift in the equilibrium, favoring the formation of the selected species [94].
The amplification of the desired compound can be efficient to enable isolation of
the molecule from the library on a preparative scale and in a high yield.
NHHN
O
O
OH
NHHN
O
O
OH
Ki = 34μM
ΔG = -6.1kcal/mol
Ki = 260μM
ΔG = -4.9kcal/mol
+
Ki = 0.0004nM
ΔG = -16.9kcal/mol
Figure 11.2 Nonadditive effect of fragment linking/merging on the binding affinity.
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The processes involved in DCC include (1) preparation of a mixture of interconverting building blocks, (2) amplification of the best binder(s) through noncovalent interactions with a template, and (3) isolat ion and characterization of the best
binder(s).
The chemical reaction that can be used in DCC must meet three sets of exacting
criteria: (1) does not disrupt the function or structure of the target, (2) must proceed
under the near physiological conditions required by the protein target (e.g., aqueous
media, physiologically compatible temperatures, and pH value), and (3) fulfils
the thermodynamic condition of reversibility. The concept of a preequilibrated
dynamic combinatorial library can be used if the reversible exchange is stopped
between generation and screening. In this case, reversible covalent reactions are
employed to reversibly form and open a chemical bond without the assistance of
the protein target. The target protein is used to preferentially select the desired
ligands [95]. The set of chemical reactions that have been successfully applied to
the construction of a dynamic combinatorial library are imine bond formation
between carbonyls and amines [96] or hydrazide [95c, 97], amide bond formation
and hydrolysis [98], disulfide bond formation between two thiols [95b, 99],
sulfide bond formation between thiols and enones [100], and alkene bond formation
by olefin metathesis [101]. The development of techniques to carefully analyze the
library composition of a DCC is necessary [102]. An approach called dynamic
combinatorial X-ray crystallography was established to visualize ligand–receptor
interactions [103].
Tethering with Extenders Tethering with extenders is a technique for extension by
tethering [70]. In this approach, a fragment that binds to the target protein is identified
first by biophysics- or bioassaybased methods. This fragment is called an extender.
The extender is modified to carry a reactive group and a protected thiol group. The
reactive functionality is used to covalently bind to the protein and to fit the fragment
into the active site. The thiol group is deprotected and used for tethering to a fragment
that binds nearby. A new molecule linking the extender and the new identified
fragment using a similar binder as that formed in the tethering study can be generated
and tested for activity.
Similar to dynamic combinatorial chemistry, tethering with extenders uses
adisulfidebondtoexploreoptimalbinders between two fragments. Tethering with
extenders has been used to identify inhibitors of cysteine aspartyl proteases
(caspase-1 [104] and caspase-3 [105]).
In Situ Click Chemistry Click chemistry was defined as a modular reaction with
wide scope, giving very high yields, stereospecific, and generating only innocuous
by-products that are easily separated. The process must include simple reaction
conditions, readily available starting materials and reagents, the use of no solvent, or
a solvent that is benign or easily removed, and simple product isolation [93]. In
contrast to dynamic combinatorial chemis try, in situ click chemistry is a kinetically
controlled TGS that irreversibly generates the products that are unable to revert to the
starting materials.
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