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Chapter
15
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Enzymatic
Catalysis
1 Catalytic Mechanisms
A. Acid–Base Catalysis
B. Covalent Catalysis
C. Metal Ion Catalysis
D. Electrostatic Catalysis
E. Catalysis through Proximity and Orientation Effects
F. Catalysis by Preferential Transition State Binding
2 Lysozyme
A. Enzyme Structure
B. Catalytic Mechanism
C. Testing the Phillips Mechanism
3 Serine Proteases
A. Kinetics and Catalytic Groups
B. X-Ray Structures
C. Catalytic Mechanism
D. Testing the Catalytic Mechanism
E. Zymogens
4 Drug Design
A. Techniques of Drug Discovery
B. Introduction to Pharmacology
C. HIV Protease and Its Inhibitors
Enzymes, as we have seen, cause rate enhancements that
are orders of magnitude greater than those of the best
chemical catalysts. Yet they operate under mild conditions
and are highly specific as to the identities of both their substrates and their products. These catalytic properties are
so remarkable that many nineteenth century scientists concluded that enzymes have characteristics that are not
shared by substances of nonliving origin. To this day, there
are few enzymes for which we understand in more than
cursory detail how they achieve their enormous rate accelerations. Nevertheless, it is now abundantly clear that
the catalytic mechanisms employed by enzymes are identical to those used by chemical catalysts. Enzymes are
simply better designed.
In this chapter we consider the nature of enzymatic
catalysis. We begin by discussing the underlying principles
of chemical catalysis as elucidated through the study of organic reaction mechanisms. We then embark on a detailed
examination of the catalytic mechanisms of several of the
best characterized enzymes: lysozyme and the serine
proteases. Their study should lead to an appreciation of
the intracacies of these remarkably efficient catalysts as
well as of the experimental methods used to elucidate their
496
properties. We end with a discussion of how drugs are discovered and tested, a process that depends heavily on the
principles of enzymology since many drug targets are
enzymes. In doing so, we consider how therapeutically
effective inhibitors of HIV-1 protease were discovered.
1 CATALYTIC MECHANISMS
Catalysis is a process that increases the rate at which a
reaction approaches equilibrium. Since, as we discussed in
Section 14-1C, the rate of a reaction is a function of its free
energy of activation (G ‡), a catalyst acts by lowering the
height of this kinetic barrier; that is, a catalyst stabilizes
the transition state with respect to the uncatalyzed reaction. There is, in most cases, nothing unique about enzymatic mechanisms of catalysis in comparison to nonenzymatic mechanisms. What apparently make enzymes such
powerful catalysts are two related properties: their specificity
of substrate binding combined with their optimal arrangement of catalytic groups. An enzyme’s arrangement of binding and catalytic groups is, of course, the product of eons
of evolution: Nature has had ample opportunity to finetune the performances of most enzymes.
The types of catalytic mechanisms that enzymes employ
have been classified as:
1. Acid–base catalysis.
2. Covalent catalysis.
3. Metal ion catalysis.
4. Electrostatic catalysis.
5. Proximity and orientation effects.
6. Preferential binding of the transition state complex.
In this section, we examine these various phenomena. In
doing so we shall frequently refer to the organic model
compounds that have been used to characterize these
catalytic mechanisms.
A. Acid–Base Catalysis
General acid catalysis is a process in which partial proton
transfer from a Brønsted acid (a species that can donate
protons; Section 2-2A) lowers the free energy of a reaction’s
transition state. For example, an uncatalyzed keto–enol
tautomerization reaction occurs quite slowly as a result of
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Section 15–1. Catalytic Mechanisms
Keto
Transition state
R
(a)
C
O
CH2
δ–
CH 2
H
H
C
O
+
H
O
H
δ+
A
δ–
C
δ–
H
H
H2O
B
A
+
OH
–
R
δ–
O
H
..
B
–
A
H
δ–
CH 2
+
+
δ+
C
H
H
CH2
H+
R
CH2
O
R
CH 2
O
H
CH2
δ–
C
A
O
δ+
CH2
C
C
R
R
(c)
R
δ–
C
R
(b)
Enol
R
O
497
C
+
H
O
H
CH2
+
δ+
+
H
H
δ+
B
+
..
7884d_c15.qxd
+
B
FIGURE 15-1 Mechanisms of keto–enol tautomerization. (a) Uncatalyzed, (b) general acid
catalyzed, and (c) general base catalyzed.
the high energy of its carbanionlike transition state (Fig.
15-1a). Proton donation to the oxygen atom (Fig. 15-1b),
however, reduces the carbanion character of the transition
state, thereby catalyzing the reaction. A reaction may also
be stimulated by general base catalysis if its rate is increased
by partial proton abstraction by a Bro/nsted base (a species
that can combine with a proton; Fig. 15-1c). Some reactions
may be simultaneously subject to both processes: a concerted general acid–base catalyzed reaction.
a. Mutarotation Is Catalyzed by Acids and by Bases
The mutarotation of glucose provides an instructive
example of acid–base catalysis. Recall that a glucose
molecule can assume either of two anomeric cyclic forms
through the intermediacy of its linear form (Section 11-1B):
H
CH2OH
O
H
OH H
H
H
OH
HO
CH2OH
O
H
OH H
H
OH
-D-Glucose
[]20
D = 18.7
H
CH2OH
OH
H
CH
OH H
HO
v
d3-D-glucose 4
dt
H
O
H
A
O
H
A
O
H
C
O
H
H
B–
-D-Glucose
-D-Glucose
H A–
O
[15.1]
where kobs is the reaction’s apparent first-order rate constant. The mutarotation rate increases with the concentrations of general acids and general bases; they are thought
to catalyze mutarotation according to the mechanism:
O
CH
H
OH
Linear form
kobs 3-D-glucose4
C
H
HO
H
OH
-D-Glucose
[]20
D = 112.2
OH
In aqueous solvents, the initial rate of mutarotation of
-D-glucose, as monitored by polarimetry (Section 4-2A),
is observed to follow the relationship:
O
H
Linear form
B
B–
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Chapter 15. Enzymatic Catalysis
This model is consistent with the observation that in
aprotic solvents such as benzene, 2,3,4,6-O-tetramethyl-D-glucose (a less polar benzene-soluble analog)
H
H3CO
CH2OCH3
O H
H
OCH3 H
OH
drolyze RNA to its component nucleotides. The isolation
of 2,3-cyclic nucleotides from RNase A digests of RNA
indicates that the enzyme mediates the following reaction
sequence:
O
–O
P
H
OCH3
2,3,4,6-O-Tetramethyl--D-glucose
5
O
CH2
4
O
O
H
Base
OH
O
–O
RNA
H
2
3
does not undergo mutarotation. Yet, the reaction is catalyzed by the addition of phenol, a weak benzene-soluble
acid, together with pyridine, a weak benzene-soluble base,
according to the rate equation:
1
H
H
P
O
CH2
O
v k3 phenol 4 3pyridine 4 3tetramethyl--D-glucose4 [15.2]
O
Base
H
H
O
OH
H
Moreover, in the presence of -pyridone, whose acid and
base groups can rapidly interconvert between two tautomeric forms and are situated so that they can simultaneously catalyze mutarotation,
–O
H
P
O
O
-Pyridone
O
N
N
O
H
H
O
H
–O
P
O
CH2
O
H
O
Base
H
H
H
O
O
C
O
H
O
O
C
H
O
+
P
–O
H
HO
CH2
O
H
2,3-Cyclic nucleotide
H
–O
v k¿ 3-pyridone 4 3tetramethyl--D-glucose4 [15.3]
H
O
the reaction follows the rate law
b. The RNase A Reaction Incorporates General
Acid–Base Catalysis
Bovine pancreatic ribonuclease A (RNase A) provides
an illuminating example of enzymatically mediated general
acid–base catalysis. This digestive enzyme functions to hy-
Base
H
Glucose
where k 7000M k. This increased rate constant indicates that -pyridone does, in fact, catalyze mutarotation
in a concerted fashion since 1M -pyridone has the same
catalytic effect as impossibly high concentrations of phenol and pyridine (e.g., 70M phenol and 100M pyridine).
Many types of biochemically significant reactions are
susceptible to acid and/or base catalysis. These include the
hydrolysis of peptides and esters, the reactions of phosphate
groups, tautomerizations, and additions to carbonyl groups.
The side chains of the amino acid residues Asp, Glu, His,
Cys, Tyr, and Lys have pK’s in or near the physiological pH
range (Table 4-1) which, we shall see, permits them to act
in the enzymatic capacity of general acid and/or base catalysts in analogy with known organic mechanisms. Indeed,
the ability of enzymes to arrange several catalytic groups
about their substrates makes concerted acid–base catalysis
a common enzymatic mechanism.
O
H2O
H+
P
OH
O
O
O
–O
P
O
CH2
O
H
O
Base
H
H
H
O
–O
P
OH
O
O–
The RNase A reaction exhibits a pH rate profile that peaks
near pH 6 (Fig. 15-2). Analysis of this curve (Section 14-4),
together with chemical derivatization and X-ray studies,
indicates that RNase A has two essential His residues, His
12 and His 119, which act in a concerted manner as general
acid and base catalysts (the structure of RNase A is
sketched in Fig. 9-2). Evidently, the RNase A reaction is a
two-step process (Fig. 15-3):
1. His 12, acting as a general base, abstracts a proton
from an RNA 2-OH group, thereby promoting its nucleo-
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Section 15–1. Catalytic Mechanisms
FIGURE 15-2 The pH dependence of V¿max K¿M in the RNase
A–catalyzed hydrolysis of cytidine-2,3-cyclic phosphate.
V¿max K¿M is given in units of M1 s1. Analysis of this curve
(Section 14-4) suggests the catalytic participation of groups with
pK’s of 5.4 and 6.4. [After del Rosario, E.J. and Hammes, G.G.,
Biochemistry 8, 1887 (1969).]
499
5
log
V'max
K'M
3
pKE1
pKE2
1
4
5
6
7
8
...
pH
...
O
P
–O
O
CH2 O
4
O
H
Base
H
O
–O
P
H
O
O
H
His 119
N
H
–O
H
P
O
N+
NH
P
H
OH
O H
–O
O
O
H
O
H
O
1
HO
N
+
H
Base
H
Base
H
H
CH2 O
H2O
His 12
CH2 O
O
NH
N
2
3
O
O
1
H
H
P
CH2 O
H
H
Base
S
–O
2′,3′-Cyclic nucleotide
O
RNA
5
H
H
O
–O
N
H
H
O
...
P
N
H
OH
O
O
...
2
...
7884d_c15.qxd
O
–O
P
O
CH2 O
H
O
Base
H
H
H
O
–O
P
O
O
H
H
N+
The bovine pancreatic RNase A–catalyzed
hydrolysis of RNA is a two-step process with the intermediate
formation of a 2,3-cyclic nucleotide.
FIGURE 15-3
O
N
H
H SN
NH
9
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Chapter 15. Enzymatic Catalysis
500
philic attack on the adjacent phosphorus atom while His
119, acting as a general acid, promotes bond scission by
protonating the leaving group.
2. The 2,3-cyclic intermediate is hydrolyzed through
what is essentially the reverse of the first step in which water replaces the leaving group. Thus His 12 acts as a general acid and His 119 as a general base to yield the hydrolyzed RNA and the enzyme in its original state.
B. Covalent Catalysis
Covalent catalysis involves rate acceleration through the
transient formation of a catalyst–substrate covalent bond.
The decarboxylation of acetoacetate, as chemically catalyzed by primary amines, is an example of such a process
(Fig. 15-4). In the first stage of this reaction, the amine nucleophilically attacks the carbonyl group of acetoacetate to
form a Schiff base (imine bond).
H
H
+
N C
H
+
N
C
O
N
C
OH
H
H
B
+
OH
–
Schiff
base
A
The protonated nitrogen atom of the covalent intermediate
then acts as an electron sink (Fig. 15-4, bottom) so as to reduce the otherwise high-energy enolate character of the transition state. The formation and decomposition of the Schiff
base occur quite rapidly, so that these steps are not rate determining in this reaction sequence.
a. Covalent Catalysis Has Both Nucelophilic and
Electrophilic Stages
As the preceding example indicates, covalent catalysis
may be conceptually decomposed into three stages:
CO2
O
CH3
C
CH2
C
CH3
C
2. The withdrawal of electrons from the reaction center by the now electrophilic catalyst.
3. The elimination of the catalyst, a reaction that is essentially the reverse of stage 1.
Reaction mechanisms are somewhat arbitrarily classified
as occurring with either nucleophilic catalysis or electrophilic catalysis depending on which of these effects provides the greater driving force for the reaction, that is,
which catalyzes its rate-determining step. The primary
amine–catalyzed decarboxylation of acetoacetate is clearly
an electrophilically catalyzed reaction since its nucleophilic
phase, Schiff base formation, is not its rate-determining
step. In other covalently catalyzed reactions, however, the
nucleophilic phase may be rate determining.
The nucleophilicity of a substance is closely related to
its basicity. Indeed, the mechanism of nucleophilic catalysis
resembles that of general base catalysis except that, instead
of abstracting a proton from the substrate, the catalyst
nucleophilically attacks it so as to form a covalent bond.
Consequently, if covalent bond formation is the ratedetermining step of a covalently catalyzed reaction, the
reaction rate tends to increase with the covalent catalyst’s
basicity (pK).
An important aspect of covalent catalysis is that the
more stable the covalent bond formed, the less facilely it
will decompose in the final steps of a reaction. A good
covalent catalyst must therefore combine the seemingly
contradictory properties of high nucleophilicity and the
ability to form a good leaving group, that is, to easily
reverse the bond formation step. Groups with high polarizabilities (highly mobile electrons), such as imidazole and
thiol functions, have these properties and hence make good
covalent catalysts.
+
H
–O
O
1. The nucleophilic reaction between the catalyst and
the substrate to form a covalent bond.
CH2
O
CH3
C
CH3
–
O
Acetoacetate
OH
Acetone
Enolate
RNH2
RNH2
OH
CH3
N
H
C
CH2
CO2
R
CH3
–
O
Schiff base
(imine)
H
N
O
C
..
R
C
CH2
H+
R
CH3
N
H
C
CH3
FIGURE 15-4 The decarboxylation of
acetoacetate. The uncatalyzed reaction
mechanism is shown at the top and the
reaction mechanism as catalyzed by
primary amines is shown at the bottom.
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Section 15–1. Catalytic Mechanisms
b. Certain Amino Acid Side Chains and Coenzymes
Can Serve as Covalent Catalysts
Enzymes commonly employ covalent catalytic mechanisms as is indicated by the large variety of covalently
linked enzyme–substrate reaction intermediates that have
been isolated. For example, the enzymatic decarboxylation
of acetoacetate proceeds, much as described above,
through Schiff base formation with an enzyme Lys residue’s
e-amino group. The covalent intermediate, in this case, has
been isolated through NaBH4 reduction of its imine bond
to an amine, thereby irreversibly inhibiting the enzyme.
Other enzyme functional groups that participate in covalent catalysis include the imidazole moiety of His, the thiol
group of Cys, the carboxyl function of Asp, and the hydroxyl group of Ser. In addition, several coenzymes, most
notably thiamine pyrophosphate (Section 17-3B) and
pyridoxal phosphate (Section 26-1A), function in association with their apoenzymes mainly as covalent catalysts.
501
The decarboxylation of dimethyloxaloacetate, as catalyzed by metal ions such as Cu2 and Ni2, is a nonenzymatic example of catalysis by a metal ion:
M n+
–O
O
O CH3
C
C
C
O
C
O–
CH3
Dimethyloxaloacetate
CO2
M n+
O– CH3
–O
C
C
C
O
CH3
C. Metal Ion Catalysis
Nearly one-third of all known enzymes require the presence
of metal ions for catalytic activity. There are two classes of
metal ion–requiring enzymes that are distinguished by the
strengths of their ion–protein interactions:
H+
–O
O
C
1. Metalloenzymes contain tightly bound metal ions,
most commonly transition metal ions such as Fe2, Fe3,
Cu2, Zn2, Mn2, or Co3.
2. Metal-activated enzymes loosely bind metal ions
from solution, usually the alkali and alkaline earth metal
ions Na, K, Mg2, or Ca2.
Metal ions participate in the catalytic process in three
major ways:
1. By binding to substrates so as to orient them properly for reaction.
2. By mediating oxidation–reduction reactions through
reversible changes in the metal ion’s oxidation state.
3. By electrostatically stabilizing or shielding negative
charges.
In this section we shall be mainly concerned with the
third aspect of metal ion catalysis. The other forms of
enzyme-mediated metal ion catalysis are considered in
later chapters in conjunction with discussions of specific
enzyme mechanisms.
a. Metal Ions Promote Catalysis through
Charge Stabilization
In many metal ion–catalyzed reactions, the metal ion
acts in much the same way as a proton to neutralize negative charge, that is, it acts as a Lewis acid. Yet metal ions
are often much more effective catalysts than protons because
metal ions can be present in high concentrations at neutral
pH’s and can have charges greater than 1. Metal ions have
therefore been dubbed “superacids.”
O
C
CH3
CH
+
Mn+
CH3
Here the metal ion (Mn), which is chelated by the dimethyloxaloacetate, electrostatically stabilizes the developing enolate ion of the transition state. This mechanism
is supported by the observation that acetoacetate, which
cannot form such a chelate, is not subject to metal ion–
catalyzed decarboxylation. Most enzymes that decarboxylate oxaloacetate require a metal ion for activity.
b. Metal Ions Promote Nucleophilic Catalysis
via Water Ionization
A metal ion’s charge makes its bound water molecules
more acidic than free H2O and therefore a source of OH
ions even below neutral pH’s. For example, the water molecule of (NH3)5Co3(H2O) ionizes according to the reaction:
1NH3 2 5Co3 1H2O2 ∆ 1NH3 2 5Co3 1OH 2 H
with a pK of 6.6, which is 9 pH units below the pK of
free H2O. The resulting metal ion–bound hydroxyl group
is a potent nucleophile.
An instructive example of this phenomenon occurs in
the catalytic mechanism of carbonic anhydrase (Section
10-1C), a widely occurring enzyme that catalyzes the
reaction:
CO2 H 2O ∆ HCO3 H Carbonic anhydrase contains an essential Zn2 ion that lies
at the bottom of an 15-Å-deep active site cleft (Fig. 8-41),
where it is tetrahedrally coordinated by three evolutionarily invariant His side chains and an O atom of either an
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Chapter 15. Enzymatic Catalysis
HCO3 ion (Fig. 15-5a) or a water molecule (Fig. 15-5b).
The enzyme has the following catalytic mechanism:
1. We begin with a water molecule bound to the protein
in the Zn2 ion’s fourth liganding position (Fig. 15-5b). This
Zn2-polarized H2O ionizes in a process facilitated through
general base catalysis by His 64 in its “in” conformation.
Although His 64 is too far away from the Zn2-bound water to directly abstract its proton, these entities are linked
by two intervening water molecules to form a hydrogen
bonded network that is thought to act as a proton shuttle.
H
H
H
N
N
H
O
H
O
H
H
O
Zn2+
Im
Im
Im
(a)
His 64
H
H
N
H
N
+
O
H
H
H
H
O–
Im
Zn2+
O
Im
Im
His 64
Im = imidazole
2. The resulting Zn2-bound OH ion nucleophilically
attacks the nearby enzymatically bound CO2, thereby converting it to HCO3 .
Im
Im
Zn
(b)
2+
Im
FIGURE 15-5 X-Ray structures of human carbonic anhydrase.
(a) Its active site in complex with bicarbonate ion. The
polypeptide is shown in ribbon form (gold) with its side chains
shown in stick form colored according to atom type (C green,
N blue, and O red). The protein-bound Zn2 ion (cyan sphere)
is tetrahedally liganded (gray bonds) by three invariant His
side chains and the HCO3 ion, which is shown in ball-and-stick
form. The HCO3 ion also interacts with the protein via van der
Waals contacts (dot surface colored according to atom type) and
a hydrogen bonded network (dashed gray lines) involving Thr
199 and Glu 106. [Based on an X-ray structure by K. K.
Kannan, Bhabha Atomic Research Center, Bombay, India.
PDBid 1HCB.] (b) The active site showing the proton shuttle
through which His 64, acting as a general base, abstracts a
proton from the Zn2-bound H2O to form an OH ion. The
polypeptide backbone is shown in ribbon form (cyan), and its
side chains and several bound solvent molecules are shown in
ball-and-stick form with C black, N blue, and O red. The
proton shuttle consists of two water molecules that form a
hydrogen bonded network (dotted white lines) that bridges the
Zn2-bound OH ion and His 64 in its “in” conformation. On
protonation, His 64 swings to the “out” conformation.
[Courtesy of David Christianson, University of Pennsylvania.]
See the Interactive Exercises
O
O–
+
C
O
H
Im
Im
O
Zn2+
O
Im
H
C
O–
H2O
Im
Im
Zn
2+
Im
O
O–
H
+
H+
+
H
O
C
O–
Im = imidazole
In doing so, the Zn2-bound OH group donates a hydrogen bond to Thr 199, which in turn donates a hydrogen
bond to Glu 106 (Fig. 15-5a). These interactions orient the
OH group with the optimal geometry (see below) for
nucleophilic attack on the substrate CO2.
3. The catalytic site is regenerated by the exchange of
the Zn2-bound HCO3 reaction product for H2O together
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Section 15–1. Catalytic Mechanisms
with the deprotonation of His 64. In the latter process, His
64 swings to its “out” conformation (Fig. 15-5b), which may
facilitate proton transfer to the bulk solvent.
503
the bimolecular reaction of imidazole with p-nitrophenylacetate,
O
c. Metal Ions Promote Reactions through
Charge Shielding
Another important enzymatic function of metal ions is
charge shielding. For example, the actual substrates of
kinases (phosphoryl-transfer enzymes utilizing ATP) are
Mg2 –ATP complexes such as
C
CH3
Adenine
Ribose
O
P
O
O–
O
P
O
P
(p-NO2Ac)
NH
O–
O
NO2
p-Nitrophenylacetate
N
O
Mg2+
O–
O
Imidazole
O–
k1
O
rather than just ATP. Here, the Mg2 ion’s role, in addition to its orienting effect, is to shield electrostatically the
negative charges of the phosphate groups. Otherwise, these
charges would tend to repel the electron pairs of attacking
nucleophiles, especially those with anionic character.
O
+
C
CH3
_
O
NO2
N
+
p-Nitrophenolate
(p-NO2O)
NH
N-Acetylimidazolium
D. Electrostatic Catalysis
The binding of substrate generally excludes water from an
enzyme’s active site. The local dielectric constant of the active site therefore resembles that in an organic solvent,
where electrostatic interactions are much stronger than
they are in aqueous solutions (Section 8-4A). The charge
distribution in a medium of low dielectric constant can
greatly influence chemical reactivity. Thus, as we have
seen, the pK’s of amino acid side chains in proteins may
vary by several units from their nominal values (Table 4-1)
because of the proximity of charged groups.
Although experimental evidence and theoretical analyses on the subject are still sparse, there are mounting indications that the charge distributions about the active sites of
enzymes are arranged so as to stabilize the transition states
of the catalyzed reactions. Such a mode of rate enhancement, which resembles the form of metal ion catalysis discussed above, is termed electrostatic catalysis. Moreover,
in several enzymes, these charge distributions apparently
serve to guide polar substrates toward their binding sites so
that the rates of these enzymatic reactions are greater than
their apparent diffusion-controlled limits (Section 14-2B).
E. Catalysis through Proximity and
Orientation Effects
Although enzymes employ catalytic mechanisms that resemble those of organic model reactions, they are far more
catalytically efficient than these models. Such efficiency
must arise from the specific physical conditions at enzyme
catalytic sites that promote the corresponding chemical reactions. The most obvious effects are proximity and
orientation: Reactants must come together with the proper
spatial relationship for a reaction to occur. For example, in
the progress of the reaction is conveniently monitored by
the appearance of the intensely yellow p-nitrophenolate
ion:
d3p-NO2
O 4
dt
k1 3 imidazole4 3p-NO2
Ac4 [15.4]
k¿1 3p-NO2
Ac4
where phenyl. Here k¿1, the pseudo-first-order rate
constant, is 0.0018 s1 when [imidazole] 1M. However,
for the intramolecular reaction
O
C
O
O
NO2
N
NH
k2
C
+
N+
–O
NO2
NH
the first-order rate constant k2 0.043 s1; that is, k2 24k¿1. Thus, when the 1M imidazole catalyst is covalently
attached to the reactant, it is 24-fold more effective than
when it is free in solution; that is, the imidazole group in
the intramolecular reaction behaves as if its concentration
is 24M. This rate enhancement has contributions from both
proximity and orientation.
a. Proximity Alone Contributes Relatively Little
to Catalysis
Let us make a rough calculation as to how the rate of
a reaction is affected purely by the proximity of its reacting groups. Following Daniel Koshland’s treatment, we
shall make several reasonable assumptions:
1. Reactant species, that is, functional groups, are
about the size of water molecules.
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Chapter 15. Enzymatic Catalysis
Thus, in the absence of other effects, this model predicts
that for the intramolecular reaction,
2. Each reactant species in solution has 12 nearestneighbor molecules, as do packed spheres of identical size.
3. Chemical reactions occur only between reactants
that are in contact.
A
4. The reactant concentration in solution is low enough
so that the probability of any reactant species being in simultaneous contact with more than one other reactant
molecule is negligible.
k1
A B ¡ A¬ B
dt
k1 3A4 3B 4 k2 3A, B4 pairs
[15.5]
where [A,B]pairs is the concentration of contacting molecules of A and B. The value of this quantity is
3A, B4 pairs 12 3A4 3B 4
[15.6]
55.5M
since there are 12 ways that A can be in contact with B,
and [A]55.5M is the fraction of sites occupied by A in water solution ([H2O] 55.5M in dilute aqueous solutions)
and hence the probability that a molecule of B will be next
to one of A. Combining Eqs. [15.5] and [15.6] yields
v k1a
55.5
b3 A, B4 pairs 4.6k1 3A, B4 pairs
12
[15.7]
R
C
R
δ–
X
X
B
b. Properly Orienting Reactants and Arresting Their
Relative Motions Can Result in Large Catalytic Rate
Enhancements
The foregoing theory is, of course, quite simple. For example, it does not take into account the relative orientations of the reacting molecules. Yet molecules are not
equally reactive in all directions as Koshland’s simple theory assumes. Rather, they react most readily only if they
have the proper relative orientation. For example, in an SN2
(bimolecular nucleophilic substitution) reaction, the incoming nucleophile optimally attacks its target C atom
along the direction opposite to that of the bond to the leaving group (Fig. 15-6). The approaches of reacting atoms
along a trajectory that deviates by as little as 10 from this
optimum direction can reduce the reaction rate by as much
as a factor of 100. In a related phenomenon, a molecule
may be maximally reactive only when it assumes a conformation that aligns its various orbitals in a way that minimizes the electronic energy of its transition state, an effect
termed stereoelectronic assistance.
obeys the second-order rate equation
d3A¬ B4
A
k2 4.6k1, which is a rather small rate enhancement.
Factors that will increase this value other than proximity
alone clearly must be considered.
Then the reaction:
v
k2
B
X–
R′′
C
R′′
R
C
R′
R′
R′′
R′
Yδ
–
Y–
Y
sp2–p hybridization at carbon
The geometry of an SN2 reaction. The attacking
nucleophile, Y, must approach the tetrahedrally coordinated
and hence sp3-hybridized C atom along the direction opposite
that of its bond to the leaving group, X, a process called
backside attack. In the transition state of the reaction, the C
atom becomes trigonal bipyramidally coordinated and hence
sp2 –p hybridized, with the p orbital (blue) forming partial bonds
to X and Y. The three sp2 orbitals form bonds to the C atom’s
FIGURE 15-6
three other substituents (R, R¿, and R– ), which have shifted
their positions into the plane perpendicular to the X¬ C¬ Y
axis (curved arrows). Any deviation from this optimal geometry
would increase the free energy of the transition state, G‡, and
hence reduce the rate of the reaction (Eq. [14.15]). The
transition state then decomposes to products in which the R,
R, and R have inverted their positions about the C atom,
which has rehybridized to sp3, and X has been released.
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Section 15–1. Catalytic Mechanisms
Another effect that we have neglected in our treatment
of proximity is that of motions of the reacting groups with
respect to one another. Yet, in the transition state complex,
the reacting groups have little relative motion. In fact, as
Thomas Bruice demonstrated, the rates of intramolecular
reactions are greatly increased by arresting a molecule’s internal motions in a way that increases the mole fraction of
the reacting groups that are in a conformation which can
enter the transition state (Table 15-1). Similarly, when an
enzyme brings two molecules together in a bimolecular
reaction, as William Jencks pointed out, not only does it
increase their proximity, but it freezes out their relative
translational and rotational motions (decreases their entropy), thereby enhancing their reactivity. Theoretical
studies by Bruice indicate that much of this rate enhancement can arise from the enzymatic binding of substrates in
a conformation that readily enters the transition state.
Enzymes, as we shall see in Sections 16-2 and 16-3, bind
substrates in a manner that both aligns and immobilizes
them so as to optimize their reactivities. The free energy
required to do so is derived from the specific binding free
energy of substrate to enzyme.
O
R1
C
O
+
R2
C
Br
O
O
R1
C
O
_
R2
+ _O
O
O
Reactantsa
Relative Rate Constant
CH3COO
Br
+
1.0
_
CH3COO
COO
Br
1 103
_
COO
COO
Br
2.3 105
_
COO
O
R
H
a
C
COOH
C
O
R
Steric
strain
R
Curved arrows indicate rotational degrees of freedom.
Source: Bruice, T.C. and Lightstone, F.C., Acc. Chem. Res. 32, 127
(1999).
as cyclopropane than for unstrained rings such as cyclohexane. In either process, the strained reactant more closely
resembles the transition state of the reaction than does the
corresponding unstrained reactant. Thus, as was first suggested by Linus Pauling and further amplified by Richard
Wolfenden and Gustav Lienhard, interactions that preferentially bind the transition state increase its concentration
and therefore proportionally increase the reaction rate.
Let us quantitate this statement by considering the kinetic consequences of preferentially binding the transition
state of an enzymatically catalyzed reaction involving a single substrate. The substrate S may react to form product
P either spontaneously or through enzymatic catalysis:
kN
H
CH2OH
8 107
_
COO
The rate enhancements effected by enzymes are often
greater than can be reasonably accounted for by the catalytic mechanisms so far discussed. However, we have not
yet considered one of the most important mechanisms of
enzymatic catalysis: the binding of the transition state to an
enzyme with greater affinity than the corresponding substrates or products. When taken together with the previously described catalytic mechanisms, preferential transition state binding rationalizes the observed rates of
enzymatic reactions.
The original concept of transition state binding proposed that enzymes mechanically strained their substrates
toward the transition state geometry through binding sites
into which undistorted substrates did not properly fit. This
so-called rack mechanism (in analogy with the medieval
torture device) was based on the extensive evidence for
the role of strain in promoting organic reactions. For
example, the rate of the reaction,
R
Br
C
COO
Br
F. Catalysis by Preferential Transition
State Binding
505
TABLE 15-1 Relative Rates of Anhydride Formation for
Esters Possessing Different Degrees of Motional Freedom
in the Reaction:
S ¡ P
kE
ES ¡ EP
+
H2O
O
Here kE and kN are the first-order rate constants for the
catalyzed and uncatalyzed reactions, respectively. The
relationships between the various states of these two reaction pathways are indicated in the following scheme:
‡
KN
KR
‡
KE
KT
ES ∆ ES‡ ¡
∆
is 315 times faster when R is CH3 rather than when it is H
because of the greater steric repulsions between the CH3
groups and the reacting groups. Similarly, ring opening reactions are considerably more facile for strained rings such
∆
‡
ES ∆ S E ¡ PE
∆
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Chapter 15. Enzymatic Catalysis
where
E+S
KR ‡
KN 3ES 4
3E4 3S4
3E4 3S‡ 4
3E4 3S 4
KT and
‡
KE 3ES‡ 4
3 E4 3S‡ 4
3ES‡ 4
∆GN
3 ES4
ES
G
are all association constants. Consequently,
‡
3S4 3ES‡ 4
KE
KT
‡
‡
KR
3S 4 3ES4
KN
∆GE
E+S
[15.8]
ES
E+P
According to transition state theory, Eqs. [14.7] and
[14.14], the rate of the uncatalyzed reaction can be expressed
kkBT
kkBT ‡
b 3S‡ 4 a
b K N 3 S4 [15.9]
vN kN 3S4 a
h
h
Similarly, the rate of the enzymatically catalyzed reaction is
EP
Reaction coordinate
FIGURE 15-7 Reaction coordinate diagrams for a hypothetical
enzymatically catalyzed reaction involving a single substrate
(blue) and the corresponding uncatalyzed reaction (red).
See the Animated Figures
kkBT
kkBT ‡
vE kE 3ES 4 a
b 3ES‡ 4 a
bKE 3 ES4 [15.10]
h
h
Therefore, combining Eqs. [15.8] to [15.10],
‡
kE
KE
KT
‡ kN
KR
KN
[15.11]
This equation indicates that the more tightly an enzyme
binds its reaction’s transition state (KT) relative to the substrate (KR), the greater the rate of the catalyzed reaction (kE)
relative to that of the uncatalyzed reaction (kN); that is,
catalysis results from the preferential binding and therefore
the stabilization of the transition state (S ‡) relative to that
of the substrate (S) (Fig. 15-7).
According to Eq. [14.15], the ratio of the rates of the
catalyzed versus the uncatalyzed reaction is expressed
kE
‡
‡
exp3 1 ¢GN ¢GE 2 RT 4
kN
a. Transition State Analogs Are Competitive Inhibitors
If an enzyme preferentially binds its transition state, then
it can be expected that transition state analogs, stable
molecules that resemble S ‡ or one of its components, are
potent competitive inhibitors of the enzyme. For example,
the reaction catalyzed by proline racemase from
Clostridium sticklandii is thought to occur via a planar transition state:
_
COO
C
C
N
A rate enhancement factor of 106 therefore requires that
an enzyme bind its transition state complex with 106-fold
higher affinity than its substrate, which corresponds to a
34.2 kJ mol1 stabilization at 25C. This is roughly the
free energy of two hydrogen bonds. Consequently, the enzymatic binding of a transition state (ES ‡) by two hydrogen bonds that cannot form in the Michaelis complex (ES)
should result in a rate enhancement of 106 based on this
effect alone.
It is commonly observed that the specificity of an enzyme is manifested by its turnover number (kcat) rather
than by its substrate-binding affinity. In other words, an
enzyme binds poor substrates, which have a low reaction
rate, as well as or even better than good ones, which have
a high reaction rate. Such enzymes apparently use a good
substrate’s intrinsic binding energy to stabilize the corresponding transition state; that is, a good substrate does not
necessarily bind to its enzyme with high affinity, but does
so on activation to the transition state.
N
H
+
H
H
[15.12]
H
+
H
_
COO
H
D-Proline
L-Proline
C
_
COO
_
N
H
Planar transition
state
Proline racemase is competitively inhibited by the planar
analogs of proline, pyrrole-2-carboxylate and -1-pyrroline-2-carboxylate,
COO–
N
H
Pyrrole-2-carboxylate
+
COO–
N
H
-1-Pyrroline-2-carboxylate
both of which bind to the enzyme with 160-fold greater
affinity than does proline. These compounds are therefore
thought to be analogs of the transition state in the proline
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Section 15–2. Lysozyme
6CH2OH
...
H
O
4
5
O
H
OH
H
H
O
1
1
CH2OH
O
H
H
H
CH2OH
O
H
H
OH
H
O
CH2OH
O
H
H
H
O
O...
H
H
H
2
3
NH
H
Lysozyme
cleavage
507
C
CH3
H
NH
O
O
C
NH
H
CH3
C
O
O
NAM
C
O
CH3
O
CH3CHCOO–
–
CH3CHCOO
NAG
NH
H
CH3
NAG
NAM
FIGURE 15-8 The alternating NAG–NAM polysaccharide component of bacterial cell walls.
The position of the lysozyme cleavage site is shown.
racemase reaction. In contrast, tetrahydrofuran-2-carboxylate,
COO–
O
H
Tetrahydrofuran-2-carboxylate
which more closely resembles the tetrahedral structure of
proline, is not nearly as good an inhibitor as these
compounds. A 160-fold increase in binding affinity corresponds, according to Eq. [15.12], to a 12.6 kJ mol1
increase in the free energy of binding. This quantity
presumably reflects the additional binding affinity that
proline racemase has for proline’s planar transition state
over that of the undistorted molecule.
Hundreds of transition state analogs for various enzymatic reactions have been reported. Some are naturally
occurring antibiotics. Others were designed to investigate
the mechanisms of particular enzymes and/or to act as specific enzymatic inhibitors for therapeutic or agricultural
use. Indeed, as we discuss in Section 15-4C, the theory that
enzymes bind transition states with higher affinity than substrates has led to a rational basis for drug design based on
the understanding of specific enzyme reaction mechanisms.
susceptible to lysozyme alone has prompted the suggestion
that this enzyme mainly helps dispose of bacteria after they
have been killed by other means.
Hen egg white (HEW) lysozyme is the most widely
studied species of lysozyme and is one of the mechanistically best understood enzymes. It is a rather small protein
(14.7 kD) whose single polypeptide chain consists of 129
amino acid residues and is internally cross-linked by four
disulfide bonds (Fig. 15-9). HEW lysozyme catalyzes the
Arg Gly Tyr Ser
Asp Asn Tyr
Leu
Gly
20
Asn
Leu
Gly
His
Trp
Arg
Ala Gln Val Asp
Trp
Lys
Gly
Cys
S
Ala
Leu
Glu
Arg
Lys Val
In the following two sections, we shall investigate the catalytic mechanisms of several well-characterized enzymes.
In doing so, we shall see how enzymes apply the catalytic
principles described in Section 15-1. You should note that
the great catalytic efficiency of enzymes arises from their simultaneous use of several of these catalytic mechanisms.
Lysozyme is an enzyme that destroys bacterial cell walls.
It does so, as we saw in Section 11-3B, by hydrolyzing the
(1S 4) glycosidic linkages from N-acetylmuramic acid
(NAM) to N-acetylglucosamine (NAG) in the alternating
NAM–NAG polysaccharide component of cell wall peptidoglycans (Fig. 15-8). It likewise hydrolyzes (1S4)-linked
poly(NAG) (chitin), a cell wall component of most fungi.
Lysozyme occurs widely in the cells and secretions of vertebrates, where it may function as a bactericidal agent.
However, the observation that few pathogenic bacteria are
Val Ala
Trp
Ser
Asn
Phe
Asn
Arg Ser
Leu
Cys
S
Gly
Pro
70 Thr
Lys
Ile
Gly
Ala
S
Leu
Leu
Cys
Ser
Asn
90
Ala Thr Ile
Gln
Ser
Asp
S
S
Ala
Thr
Asn
Asn
Ser
Lys
Thr 40
Arg
Cys 80
Ile
Asn
Asn
Pro
Val
Ser
Trp
Asn
Met
Ser 100
Val
Glu
110
Ala
Gly
Ala
Asn
Phe
Asp
Asp
Ala
Arg
Gly
2 LYSOZYME
Ala
Lys
COO–
Arg
H3N+
Cys 30
Arg
Leu
S
Gly
Phe
S
Cys
129
Cys
1
S
Lys
Arg
Ala
Ala
10
Gly
120
Ile
Met
Val
Thr
Asp
Arg
Cys
Asn
Trp
Thr
Trp
Asp
Arg
Gly
Ser 60
50 Ser
Asn
Thr
Ile
Gln
Leu Ile Gly
Asp
Tyr
FIGURE 15-9 Primary structure of HEW lysozyme. The amino
acid residues that line the substrate-binding pocket are shown
in dark purple.
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508
Chapter 15. Enzymatic Catalysis
hydrolysis of its substrate at a rate that is 108-fold greater
than that of the uncatalyzed reaction.
A. Enzyme Structure
The elucidation of an enzyme’s mechanism of action requires a knowledge of the structure of its enzyme–substrate
complex. This is because, even if the active site residues
have been identified through chemical and physical means,
their three-dimensional arrangements relative to the substrate as well as to each other must be known for an
understanding of how the enzyme works. However, an enzyme binds its good substrates only transiently before it
catalyzes a reaction and releases the products. Consequently, most of our knowledge of enzyme–substrate
complexes derives from X-ray studies of enzymes in complex with inhibitors or poor substrates that remain stably
bound to the enzyme for the several hours that are usually
required to measure a protein crystal’s X-ray diffraction
intensities (although techniques for measuring X-ray intensities in less than 1 s have been developed). The large
solvent-filled channels that occupy much of the volume of
most protein crystals (Section 8-3A) often permit the formation of enzyme–inhibitor complexes by the diffusion of
inhibitor molecules into crystals of the native protein.
The X-ray structure of HEW lysozyme, which was elucidated by David Phillips in 1965, was the second structure
of a protein and the first of an enzyme to be determined
at high resolution. The protein molecule is roughly ellipsoidal in shape with dimensions 30 30 45 Å (Fig.
15-10). Its most striking feature is a prominent cleft, the
substrate-binding site, that traverses one face of the molecule. The polypeptide chain forms five helical segments as
well as a three-stranded antiparallel sheet that comprises
much of one wall of the binding cleft (Fig. 15-10b). As expected, most of the nonpolar side chains are in the interior
of the molecule, out of contact with the aqueous solvent.
ture of the (NAG)3 –lysozyme complex reveals that
(NAG)3 is bound on the right side of the enzymatic binding cleft as drawn in Fig. 15-10a for substrate residues A,
B, and C. This inhibitor associates with the enzyme through
strong hydrogen bonding interactions, some of which involve the acetamido groups of residues A and C, as well
as through close-fitting hydrophobic contacts. In an example of induced-fit ligand binding (Section 10-4C), there
is a slight (1 Å) closure of lysozyme’s binding cleft on
binding (NAG)3.
b. Lysozyme’s Catalytic Site Was Identified through
Model Building
(NAG)3 takes several weeks to hydrolyze under the influence of lysozyme. It is therefore presumed that the complex revealed by X-ray analysis is unproductive; that is, the
enzyme’s catalytic site occurs at neither the A¬ B nor the
B¬ C bonds. [Presumably, the rare occasions when
(NAG)3 hydrolyzes occur when it binds productively at the
catalytic site.]
In order to locate lysozyme’s catalytic site, Phillips used
model building to investigate how a larger substrate could
bind to the enzyme. Lysozyme’s active site cleft is long
enough to accommodate (NAG)6, which the enzyme
rapidly hydrolyzes (Table 15-2). However, the fourth NAG
residue (residue D in Fig. 15-10a) appeared unable to bind
to the enzyme because its C6 and O6 atoms too closely
contact Glu 35, Trp 108, and the acetamido group of NAG
residue C. This steric interference could be relieved by distorting the glucose ring from its normal chair conforma-
(Opposite) X-Ray structure of HEW lysozyme.
(a) The polypeptide chain is shown with a bound (NAG)6
substrate (green). The positions of the backbone C atoms are
indicated together with those of the side chains that line the
substrate-binding site and form disulfide bonds. The substrate’s
sugar rings are designated A, at its nonreducing end (right),
through F, at its reducing end (left). Lysozyme catalyzes the
hydrolysis of the glycosidic bond between residues D and E.
Rings A, B, and C are observed in the X-ray structure of the
complex of (NAG)3 with lysozyme; the positions of rings D, E,
and F were inferred from model building studies. [Illustration,
Irving Geis/Geis Archives Trust. Copyright Howard Hughes
Medical Institute. Reproduced with permission.] (b) A ribbon
diagram of lysozyme highlighting the protein’s secondary
structure and indicating the positions of its catalytically
important side chains, Glu 35 and Asp 52 (red). (c) A
computer-generated model showing the protein’s molecular
envelope ( purple) and C backbone (blue). The side chains of
the catalytic residues, Asp 52 (above) and Glu 35 (below), are
colored yellow. Note the enzyme’s prominent substrate-binding
cleft. [Courtesy of Arthur Olson, The Scripps Research
Institute, La Jolla, California.] Parts a, b, and c have
See the Interactive
approximately the same orientation.
FIGURE 15-10
a. The Nature of the Binding Site
NAG oligosaccharides of less than five residues are but
very slowly hydrolyzed by HEW lysozyme (Table 15-2) although these substrate analogs bind to the enzyme’s active
site and are thus its competitive inhibitors. The X-ray struc-
Rates of HEW Lysozyme-Catalyzed
Hydrolysis of Selected Oligosaccharide Substrate Analogs
TABLE 15-2
Compound
kcat (s1)
(NAG)2
2.5 108
(NAG)3
8.3 106
(NAG)4
6.6 105
(NAG)5
0.033
(NAG)6
0.25
(NAG–NAM)3
0.5
Source: Imoto, T., Johnson, L.N., North, A.C.T., Phillips, D.C., and
Rupley, J.A., in Boyer, P.D. (Ed.), The Enzymes (3rd ed.), Vol. 7, p. 842,
Academic Press (1972).
Exercises and Kinemage Exercise 9
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Section 15–2. Lysozyme
(a)
(b)
(c)
N
Asp 52
Glu 35
C
509
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Chapter 15. Enzymatic Catalysis
510
OH
tion to that of a half-chair (Fig. 15-11). This distortion,
which renders atoms C1, C2, C5, and O5 of residue D coplanar, moves the ¬ C6H2OH group from its normal equatorial position to an axial position where it makes no close
contacts and can hydrogen bond to the backbone carbonyl
group of Gln 57 and the amido group of Val 109 (Fig. 1512). Continuing the model building, Phillips found that
residues E and F apparently bind to the enzyme without
distortion and with a number of favorable hydrogen bonding and van der Waals contacts.
We are almost in a position to identify lysozyme’s catalytic site. In the enzyme’s natural substrate, every second
residue is an NAM. Model building, however, indicated
that its lactyl side chain cannot be accommodated in the
binding subsites of either residues C or E. Hence, the NAM
CH2OH
HO
O
A
NAG
–O
C
H3C
O
N
C
Asp 101
O
H
O
R
H
O
O
CH2
B
H
NAM
O
N
C
H3C
O
O
H
NAG NAM
A
B
NAG NAM
C
D
NAG NAM
E
F
(
)
reducing
end
Trp
62
O
CH2
N
C
Asn
59 N
N
R
D ring in
half-chair
conformation
O
Val
109
CH2O
D
O
C
NAM H O
O
O
– C
C
H
O
Gln
57
Asp 52
O
Lysozyme cuts
N
H
O
C
C
O
H
NH2
O
E
NAG
C
C
Glu 35
CH2OH O
O
O
N
O
H3C
Asn
44
Ala
107
C
CH3
H
O5
C5
O
H
C
O
H
NH2
C4
NAG
O
Gln
57
Trp
63
N
O
H
H
residues must bind to the enzyme in subsites B, D, and F.
The observation that lysozyme hydrolyzes (1S4) linkages from NAM to NAG implies that bond cleavage occurs
either between residues B and C or between residues D
and E. Since (NAG)3 is stably bound to but not cleaved
by the enzyme while spanning subsites B and C, the probable cleavage site is between residues D and E. This conclusion is supported by John Rupley’s observation that
lysozyme nearly quantitatively hydrolyzes (NAG)6
H
O
N
H3C
H
C2
C3
Glu
C
35
C1
O
O
Chair conformation
R
H
CH2
F
C
C4
H3C
O
C3
C
Asn
37
H2N
NAM
O
O
N
H
O5
C
O
O
O
C5
Phe
34
O
H2N
+
H2N
NH
H
C2
C1
Half-chair conformation
FIGURE 15-11 Chair and half-chair conformations. Hexose
rings normally assume the chair conformation. It is postulated,
however, that binding by lysozyme distorts the D-ring into the
half-chair conformation such that atoms C1, C2, C5, and O5 are
See the Animated Figures
coplanar.
Interactions of lysozyme with its substrate. The
view is into the binding cleft with the heavier edges of the rings
facing the outside of the enzyme and the lighter ones against
the bottom of the cleft. [Illustration, Irving Geis/Geis Archives
Trust. Copyright Howard Hughes Medical Institute. Reproduced
with permission. Based on an X-ray structure by David Phillips,
See Kinemage
Oxford University, U.K. PDBid 4LYZ.]
FIGURE 15-12
Exercise 9
Arg
114
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Section 15–2. Lysozyme
between the second and third residues from its reducing
terminus (the end with a free C1¬OH), just as is expected
if the enzyme has six saccharide-binding subsites and
cleaves its bound substrate between residues D and E.
The bond that lysozyme cleaves was identified by carrying out the lysozyme-catalyzed hydrolysis of (NAG)3 in
H218O. The resulting product had 18O bonded to the C1
atom of its newly liberated reducing terminus, thereby
demonstrating that bond cleavage occurs between C1 and
the bridge oxygen O1:
OR
H
C
511
OR
O
R
+
H
+
H
R
C
+
O
R
H
Acetal
R
ROH
R
H
R
+
O
O
C
C
+
R
H
R
Resonance-stabilized
carbocation (oxonium ion)
H
O
C1
H
O1
C
4
NAc
18
OR
OH
CH2OH
H
OH
C
+
H
NAc
C
+
H
H
H
OH
HO
H
C
OH
R
lysozyme
18
H
H
H
H2 O
O
H2O
CH2OH
Hemiacetal
Mechanism of the nonenzymatic acid-catalyzed
hydrolysis of an acetal to a hemiacetal. The reaction involves
the protonation of one of the acetal’s oxygen atoms followed by
cleavage of its C¬ O bond to form an alcohol (ROH) and a
resonance-stabilized carbocation (oxonium ion). The addition
of water to the oxonium ion forms the hemiacetal and
regenerates the H catalyst. Note that the oxonium ion’s C, O,
H, R, and R atoms all lie in the same plane.
FIGURE 15-13
Thus, lysozyme catalyzes the hydrolysis of the C1 ¬ O1
bond of a bound substrate’s D residue. Moreover, this
reaction occurs with retention of configuration, so that the
D-ring product remains the anomer.
B. Catalytic Mechanism
It remains to identify lysozyme’s catalytic groups. The
reaction catalyzed by lysozyme, the hydrolysis of a glycoside, is the conversion of an acetal to a hemiacetal.
Nonenzymatic acetal hydrolysis is an acid-catalyzed reaction that involves the protonation of a reactant oxygen
atom followed by cleavage of its C ¬ O bond (Fig. 15-13).
This results in the formation of a resonance-stabilized carbocation that is called an oxonium ion. To attain maximum
orbital overlap, and thus resonance stabilization, the
oxonium ion’s R and R groups must be coplanar with its
C, O, and H atoms (stereoelectronic assistance). The oxonium ion then adds water to yield the hemiacetal and regenerate the acid catalyst. In searching for catalytic groups
on an enzyme that mediates acetal hydrolysis, we should
therefore seek a potential acid catalyst and possibly a
group that could further stabilize an oxonium ion
intermediate.
a. Glu 35 and Asp 52 Are Lysozyme’s
Catalytic Residues
The only functional groups in the immediate vicinity of
lysozyme’s reaction center that have the required catalytic
properties are the side chains of Glu 35 and Asp 52,
residues that are invariant in the family of lysozymes of
which HEW lysozyme is the prototype. These side chains,
which are disposed to either side of the (1S 4) glycosidic
linkage to be cleaved (Fig. 15-10), have markedly different
environments. Asp 52 is surrounded by several conserved
polar residues with which it forms a complex hydrogen
bonded network. Asp 52 is therefore predicted to have a
normal pK; that is, it should be unprotonated and hence
negatively charged throughout the 3 to 8 pH range in which
lysozyme is catalytically active. In contrast, the carboxyl
group of Glu 35 is nestled in a predominantly nonpolar
pocket, where, as we discussed in Section 15-1D, it is likely
to remain protonated at unusually high pH’s for carboxyl
groups. Indeed, neutron diffraction studies, which provide
similar information to X-ray diffraction studies but also reveal the positions of hydrogen atoms, indicate that Glu 35
is protonated at physiological pH’s. The closest approaches
in the X-ray structures between the carboxyl O atoms of
both Asp 52 and Glu 35 and the C1 ¬ O1 bond of NAG
residue D are 3 Å, which makes them the prime candidates for electrostatic and acid catalysts, respectively.
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Chapter 15. Enzymatic Catalysis
b. The Phillips Mechanism
With much of the foregoing information, Phillips postulated the following enzymatic mechanism for lysozyme
(Fig. 15-14):
1. Lysozyme attaches to a bacterial cell wall by binding
to a hexasaccharide unit. In the process, residue D is distorted toward the half-chair conformation in response to
the unfavorable contacts that its ¬ C6H2OH group would
otherwise make with the protein.
Lysozyme,
main chain
Asp 52
O– NAc
2. Glu 35 transfers its proton to the O1 of the D-ring,
the only polar group in its vicinity (general acid catalysis).
The C1 ¬ O1 bond is thereby cleaved, generating a resonance-stabilized oxonium ion at C1.
3. The ionized carboxyl group of Asp 52 acts to stabilize
the developing oxonium ion through charge–charge interactions (electrostatic catalysis). This carboxylate group
apparently cannot form a covalent bond with the substrate
because the observed 3 Å distance between C1 and a carboxyl O atom of Asp 52 is much greater than the 1.4 Å
length of a C¬ O covalent bond [i.e., the reaction appears
to occur via an SN1 (unimolecular nucleophilic substitution) mechanism to yield an oxonium ion, not via a
mechanism involving the transient formation of a C ¬ O
covalent bond to the enzyme; but see Section 15-2C]. The
bond cleavage reaction is facilitated by the strain in the
D-ring that distorts it to the planar half-chair conformation. This is a result of the oxonium ion’s required planarity; that is, the initial binding conformation of the D-ring
resembles that of the reaction’s transition state (transition
state binding catalysis; Fig. 15-15).
4. At this point, the enzyme releases the hydrolyzed
E-ring with its attached polysaccharide (the leaving group),
yielding a cationic, noncovalent, glycosyl–enzyme intermediate. This oxonium ion subsequently adds H2O from
solution in a reversal of the preceding steps to form product and to reprotonate Glu 35. The reaction’s retention of
configuration is dictated by the shielding of one of the oxonium ion’s faces by the enzymatic cleft. The enzyme then
releases the D-ring product with its attached saccharide,
thereby completing the catalytic cycle.
D
O+
CH2OH
O–
CH2OH
Glu 35
H+
Lysozyme,
main chain
H2O
The Phillips mechanism for the lysozyme
reaction. The cleavage of the glycosidic bond between the
substrate D- and E-rings occurs through protonation of the
bridge oxygen atom by Glu 35. The resulting D-ring oxonium
ion is stabilized by the proximity of the Asp 52 carboxylate
group and the enzyme-induced distortion of the D-ring. Once
the E-ring is released, H2O from solution provides both an
OH that combines with the oxonium ion and an H that
reprotonates Glu 35. NAc represents the N-acetylamino
See Kinemage
substituent at C2 of each glucose ring.
FIGURE 15-14
Exercise 9 and the Animated Figures
CH2OH
O
H
C
+
H
OR
H
H
NHCOCH3
H
.
O
H
..
.
Glu 35. The mutagenesis of Glu 35 to Gln yields a protein
with no detectable catalytic activity (0.1% of wild type),
..
a. Identification of the Catalytic Residues
Lysozyme’s catalytically important groups have been
experimentally identified through site-directed mutagenesis (Section 5-5G) and the use of group-specific reagents:
OH–
NAc
C. Testing the Phillips Mechanism
The Phillips mechanism was formulated largely on the basis of structural investigations of lysozyme and a knowledge of the mechanism of nonenzymatic acetal hydrolysis.
A variety of evidence has since been gathered that bears
on the validity of this mechanism. In the remainder of this
section, we discuss the highlights of these studies to illustrate how scientific models evolve.
H+
E
O
CH2OH
+
O
H
C H
OR
H
H
NHCOCH3
CH3
R = H (NAG) or
CH (NAM)
_
COO
The D-ring oxonium ion intermediate in the
Phillips mechanism is stabilized by resonance. This requires
that atoms C1, C2, C5, and O5 be coplanar (shading); that is,
the hexose ring must assume the half-chair conformation.
FIGURE 15-15
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Section 15–2. Lysozyme
although it has only a 1.5-fold decrease in substrate affinity. Glu 35 must therefore be essential for lysozyme’s
catalytic activity.
Asp 52. The mutagenesis of Asp 52 to Asn, which has a
polarity comparable to that of Asp but lacks its negative
charge, yields an enzyme with no more than 5% of wildtype lysozyme’s catalytic activity even though this mutation
causes an 2-fold increase in the enzyme’s affinity for
substrate. Asp 52 is therefore important for enzymatic
activity.
Noninvolvement of Other Amino Acid Residues.
Lysozyme’s other carboxyl groups besides Glu 35 and Asp
52 do not participate in the catalytic process, as was
demonstrated by reacting lysozyme with carboxyl-specific
reagents in the presence of substrate. This treatment yields
an almost fully active enzyme in which all carboxyl groups
but Glu 35 and Asp 52 are derivatized. Other group-specific
reagents that modify, for instance, His, Lys, Met, or Tyr
residues but induce no major protein structure disruptions
cause little change in lysozyme’s catalytic efficiency.
b. Role of Strain
Many of the mechanistic investigations of lysozyme have
had the elusive goal of establishing the catalytic role of
strain. Not all of these studies, as we shall see, have supported the Phillips mechanism, thereby stimulating a series of investigations that have only recently settled this
issue.
Measurements of the binding equilibria of various
oligosaccharides to lysozyme indicate that all saccharide
residues except that binding to the D subsite contribute energetically toward the binding of substrate to lysozyme;
binding NAM in the D subsite requires a free energy input
of 12 kJ mol1 (Table 15-3). The Phillips mechanism
explains this observation as being indicative of the energy
penalty of straining the D-ring from its preferred chair
conformation toward the half-chair form.
As we have discussed in Section 15-1F, an enzyme that
catalyzes a reaction by the preferential binding of its transition state has a greater binding affinity for an inhibitor
that has the transition state geometry (transition state analog) than it does for its substrate. The -lactone analog of
(NAG)4 (Fig. 15-16) is a transition state analog of lysozyme
TABLE 15-3
Binding Free Energies of HEW Lysozyme
Subsites
Site
Bound
Saccharide
Binding
Free Energy
(kJ mol1)
A
NAG
7.5
B
NAM
12.3
C
NAG
23.8
D
NAM
12.1
E
NAG
7.1
F
NAM
7.1
Source: Chipman, D.M. and Sharon, N., Science 165, 459 (1969).
H
O
(NAG)3
CH2OH
O
H
H
O
OH
H
H
NHCOCH3
O
(NAG)3
CH2OH
+
O
H
513
–
O
OH
H
H
NHCOCH3
The -lactone analog of (NAG)4. Its C1, O1,
C2, C5, and O5 atoms are coplanar (shading) because of
resonance, as is the D-ring in the reaction intermediate of the
Phillips mechanism (compare with Fig. 15-15).
FIGURE 15-16
since this compound’s lactone ring has the half-chair conformation that geometrically resembles the proposed oxonium ion transition state of the substrate’s D-ring. X-Ray
studies indicate, in accordance with prediction, that this inhibitor binds to lysozyme’s A¬ B¬ C ¬ D subsites such
that the lactone ring occupies the D subsite in a halfchairlike conformation.
Despite the foregoing, the role of substrate distortion in
lysozyme catalysis had been questioned. Theoretical studies by Michael Levitt and Arieh Warshel on substrate binding by lysozyme suggested that the protein is too flexible
to mechanically distort the D-ring of a bound substrate.
Rather, these calculations implied that transition state stabilization occurs through the displacement by substrate of
several tightly bound water molecules from the D subsite.
The resulting desolvation of the Asp 52 carboxylate group
would significantly enhance its capacity to electrostatically
stabilize the transition state oxonium ion. This study therefore concluded that “electrostatic strain” rather than steric
strain is the more important factor in stabilizing lysozyme’s
transition state.
In an effort to obtain further experimental information
bearing on the Phillips strain mechanism, Nathan Sharon
and David Chipman determined the D subsite–binding
affinities of several saccharides by comparing the
lysozyme-binding affinities of various substrate analogs.
The NAG lactone inhibitor binds to the D subsite with 9.2
kJ mol1 greater affinity than does NAG. This quantity
corresponds, according to Eq. [14.15], to no more than an
40-fold rate enhancement of the lysozyme reaction as a
result of strain (recall that the difference in binding energy
between a transition state analog and a substrate is indicative of the enzyme’s rate enhancement arising from the
preferential binding of the transition state complex). Such
an enhancement is hardly a major portion of lysozyme’s
108-fold rate enhancement (accounting for only 20%
of the reaction’s ¢ ¢G‡cat ; Section 14-1C). Moreover, an
N-acetylxylosamine (XylNAc) residue,
H
O
H
O
H
OH
OH
H
H
H
NHCOCH3
N -Acetylxylosamine residue
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Chapter 15. Enzymatic Catalysis
which lacks the sterically hindered ¬ C6H2OH group of
NAM and NAG, has only marginally greater binding affinity for the D subsite (3.8 kJ mol1) than does NAG
(2.5 kJ mol1). Yet recall that the Phillips mechanism
postulates that it is the unfavorable contacts made by this
¬ C6H 2OH group that promotes D-ring distortion.
Nevertheless, lysozyme does not hydrolyze saccharides
with XylNAc in the D subsite.
The apparent inconsistencies among the foregoing experimental observations were largely rationalized by
Michael James’ highly accurate (1.5-Å resolution) X-ray
crystal structure determination of lysozyme in complex
with NAM–NAG–NAM. This trisaccharide binds, as expected, to the B, C, and D subsites of lysozyme. The NAM
in the D subsite, in agreement with the Phillips mechanism,
is distorted to the half-chair conformation with its
¬ C6H2OH group in a nearly axial position due to steric
clashes that would otherwise occur with the acetamido
group of the C subsite NAG (although, contrary to the original Phillips mechanism, Glu 35 and Trp 108 are too far
away from the ¬ C6H2OH group to contribute to this distortion). This strained conformation is stabilized by a
strong hydrogen bond between the D-ring O6 and the
backbone NH of Val 109 (transition state stabilization).
Indeed, the mutation of Val 109 to Pro, which lacks the
NH group to make such a hydrogen bond, inactivates the
enzyme. Lysozyme’s lack of hydrolytic activity when
XylNAc occupies its D subsite is likewise explained by the
absence of this hydrogen bond and the consequent lesser
stability of the XylNAc ring’s half-chair transition state.
The unexpectedly small free energy differences in binding NAG, NAG lactone, and XylNAc to the D subsite are
explained by the observation that undistorted NAG and
XylNAc can be modeled into the D subsite as it occurs in
the X-ray structure of the lysozyme NAM–NAG–NAM
complex. NAM’s bulky lactyl side chain prevents it from
binding to the D subsite in this manner.
c. The Lysoyme Reaction Proceeds via a
Covalent Intermediate
Alternatives to the Phillips mechanism postulate that
either (1) the carboxyl group of Asp 52 displaces the leaving group to form a covalent bond to C1, thereby yielding
a covalent glycosyl–enzyme ester intermediate that is subsequently displaced by water to yield product (a doubledisplacement mechanism); or (2) water directly displaces
the leaving group (a single-displacement mechanism). A
single-displacement mechanism would result in inversion
of configuration between substrate and product and thus
can be ruled out. A double-displacement mechanism
would account for the observed retention of configuration
in the lysozyme reaction (as does the Phillips mechanism).
However, it is at odds with the observation that the distance between C1 in a D subsite–bound saccharide and a
carboxyl O of Asp 52 (which participates in a network of
hydrogen bonds that apparently hold this side chain in its
position) are too long to form a covalent bond (minimally
2.3 Å in the NAM–NAG–NAM complex without significantly disrupting the protein structure vs 1.4 Å for a
C¬ O single bond). Indeed, no such covalent bond had
been observed in any of the numerous X-ray structures
containing hen egg white (HEW) lysozyme.
Despite the foregoing, all other -glycosidases of known
structure that cleave glycosidic linkages with net retention
of configuration at the anomeric carbon (as does HEW
lysozyme) have been shown to do so via a covalent
glycosyl–enzyme intermediate. The active sites of these socalled retaining -glycosidases structurally resemble that
of HEW lysozyme. Moreover, there is no direct evidence
indicative of the existence of a long-lived oxonium ion at
the active site of any retaining -glycosidase, including
HEW lysozyme (the lifetime of a glucosyl oxonium ion in
water is 1012 s, a time only slightly longer than that of
a bond vibration). Consequently, there had been a growing
suspicion that the HEW lysozyme reaction also proceeds
via a covalent intermediate, one between the D-ring’s
anomeric carbon (C1) and the side chain carboxyl group
of Asp 52 to form an ester linkage:
H
CH2OH
O
H
OR
H
O
O
C
CH2
Asp 52
H
O
H
NHCOCH3
This intermediate presumably reacts with H2O in what is
essentially the reverse of the reaction leading to its formation, thereby yielding the reaction’s second product
(a double-displacement mechanism). In this mechanism,
the oxonium ion is proposed to be the transition state on
the way to forming the covalent intermediate, rather than
being an intermediate itself.
If, in fact, HEW lysozyme follows this mechanism, the
reason that its covalent intermediate had never been observed is that its rate of breakdown must be much faster
than its rate of formation. Hence, if this intermediate is to
be experimentally observed, its rate of formation must be
made significantly greater than its rate of breakdown. To
do so, Stephen Withers capitalized on three phenomena.
First, if, as postulated, the reaction goes through an oxonium ion transition state, all steps involving its formation
should be slowed by the electron withdrawing effects of
substituting F (the most electronegative element) at C2 of
the D-ring. Second, mutating Glu 35 to Gln (E35Q) removes the general acid–base that catalyzes the reaction,
further slowing all steps involving the oxonium ion transition state. Third, substituting an additional F atom at C1
of the D-ring accelerates the formation of the intermediate because this F is a good leaving group. Making all three
of these changes should increase the rate of formation
of the proposed covalent intermediate relative to its
breakdown and hence should result in its accumulation.
Withers therefore incubated E35Q HEW lysozyme with
NAG-(1S 4)-2-deoxy-2-fluoro--D-glucopyranosyl fluoride (NAG2FGlcF):
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Section 15–3. Serine Proteases
H
CH2OH
O
H
OH H
HO
H
O
CH2OH
O
H
OH H
H
H
NHCOCH3
F
H
H
F
NAG2FGlcF
Electrospray ionization mass spectrometry (ESI-MS; Section 7-1J) of this reaction mixture revealed a sharp peak
at 14,683 D, consistent with the formation of the proposed
covalent intermediate, but no significant peak at or near
the 14,314-D molecular mass of the mutant enzyme alone.
The X-ray structure of this covalent complex unambiguously reveals the expected 1.4-Å-long covalent bond
between C1 of the D-ring NAG and a side chain carboxyl
O of Asp 52 (Fig. 15-17). This D-ring NAG adopts an undis-
515
torted chair conformation, thus indicating that it is a reaction intermediate rather than an approximation of the transition state. The superposition of this covalent complex with
that of the above described complex of NAM–NAG–NAM
with wild-type HEW lysozyme reveals how this covalent
bond forms (Fig. 15-17). The shortening of the 3.2-Å distance between the D-ring NAG C1 and the Asp 52 O in
the NAM–NAG–NAM complex to 1.4 Å in the covalent
complex is almost entirely a consequence of the relaxation
of the D-ring from the half-chair to the chair conformation
combined with an 45 rotation of the Asp 52 side chain
about its C ¬ C bond; the positions of the D-ring O4 and
O6 atoms are essentially unchanged. Hence, over 35 years
after its formulation, it was shown that the Phillips mechanism must be altered to take into account the transient formation of this covalent glycosyl–enzyme ester intermediate
(covalent catalysis). Keep in mind, however, that in order
to form this covalent linkage, the D-ring must pass through
an oxonium-like transition state, which requires it to transiently assume the half-chair conformation.
3 SERINE PROTEASES
Our next example of enzymatic mechanisms is a diverse
group of proteolytic enzymes known as the serine proteases
(Table 15-4). These enzymes are so named because they
have a common catalytic mechanism characterized by the
possession of a peculiarly reactive Ser residue that is essential for their enzymatic activity. The serine proteases are the
most thoroughly understood family of enzymes as a result
of their extensive examination over a nearly 50-year period
by kinetic, chemical, physical, and genetic techniques. In this
section, we mainly study the best characterized serine proteases, chymotrypsin, trypsin, and elastase. We also consider
how these three enzymes, which are synthesized in inactive
forms, are physiologically activated.
A. Kinetics and Catalytic Groups
The HEW lysozyme covalent intermediate. The
substrate C- and D-rings and Asp 52 are shown in the
superposition of the X-ray structures of the covalent complex
formed by reacting E35Q lysozyme with NAG2FGlcF (C green,
N blue, O red, and F magenta) and the noncovalent complex of
wild-type lysozyme with NAM–NAG–NAM (C yellow, N blue,
and O red). Note that the covalent bond between Asp 52 and
C1 of the D-ring forms when the D-ring in the noncovalent
complex relaxes from its distorted half-chair conformation to an
undistorted chair conformation and that the side chain of Asp
52 undergoes an 45 rotation about its C¬ C bond. [Based
on X-ray structures by David Vocadlo and Stephen Withers,
University of British Columbia, Vancouver, Canada; and
Michael James, University of Alberta, Edmonton, Canada.
PDBids 1H6M and 9LYZ.]
FIGURE 15-17
Chymotrypsin, trypsin, and elastase are digestive enzymes
that are synthesized by the pancreatic acinar cells (Fig. 1-10c)
and secreted, via the pancreatic duct, into the duodenum (the
small intestine’s upper loop). All of these enzymes catalyze
the hydrolysis of peptide (amide) bonds but with different
specificities for the side chains flanking the scissile (to be
cleaved) peptide bond (recall that chymotrypsin is specific
for a bulky hydrophobic residue preceding the scissile peptide bond, trypsin is specific for a positively charged residue,
and elastase is specific for a small neutral residue; Table 72). Together, they form a potent digestive team.
a. Ester Hydrolysis as a Kinetic Model
That chymotrypsin can act as an esterase as well as a
protease is not particularly surprising since the chemical
mechanisms of ester and amide hydrolysis are almost identical. The study of chymotrypsin’s esterase activity has led
to important insights concerning this enzyme’s catalytic
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Chapter 15. Enzymatic Catalysis
TABLE 15-4
A Selection of Serine Proteases
Enzyme
Source
Function
Trypsin
Pancreas
Digestion of proteins
Chymotrypsin
Pancreas
Digestion of proteins
Elastase
Pancreas
Digestion of proteins
Thrombin
Vertebrate serum
Blood clotting
Plasmin
Vertebrate serum
Dissolution of blood clots
Kallikrein
Blood and tissues
Control of blood flow
Complement C1
Serum
Cell lysis in the immune response
Acrosomal protease
Sperm acrosome
Penetration of ovum
Lysosomal protease
Animal cells
Cell protein turnover
Cocoonase
Moth larvae
Dissolution of cocoon after metamorphosis
-Lytic protease
Bacillus sorangium
Possibly digestion
Proteases A and B
Streptomyces griseus
Possibly digestion
Subtilisin
Bacillus subtilis
Possibly digestion
Source: Stroud, R.M., Sci. Am. 231(1), 86 (1974).
Time course of p-nitrophenylacetate hydrolysis
as catalyzed by two different concentrations of chymotrypsin.
The enzyme rapidly binds substrate and releases the first
product, p-nitrophenolate ion, but the second product, acetate
ion, is released more slowly. Consequently, the rate of
p-nitrophenolate generation begins rapidly (burst phase) but
slows as acyl–enzyme complex accumulates until the rate of
p-nitrophenolate generation approaches that of acetate release
(steady state). The extrapolation of the steady state curve to
zero time (dashed lines) indicates the initial concentration of
active enzyme. [After Hartley, B.S. and Kilby, B.A., Biochem. J.
56, 294 (1954).]
4
FIGURE 15-18
[ p-Nitrophenolate] (mM)
Burst
phase
Steady state
phase
L–1
g⋅m
0.8 m
3
2
–1
0.4 mg ⋅ mL
1
0
2
4
6
Time (min)
mechanism. Kinetic measurements by Brian Hartley of the
chymotrypsin-catalyzed hydrolysis of p-nitrophenylacetate
O
CH3
C
NO2
O
8
10
12
p-nitrophenolate ion forming a covalent acyl–enzyme intermediate that (2) is slowly hydrolyzed to release acetate:
O
p-Nitrophenylacetate
H2O
chymotrypsin
2H+
CH3
C
O
NO2
p-Nitrophenylacetate
+
Chymotrypsin
O
CH3
C
Acetate
O–
+
–O
NO2
–O
fast
O
indicated that the reaction occurs in two phases (Fig. 15-18):
1. The “burst phase,” in which the highly colored
p-nitrophenolate ion is rapidly formed in amounts
stoichiometric with the quantity of active enzyme present.
2. The “steady-state phase,” in which p-nitrophenolate
is generated at a reduced but constant rate that is independent of substrate concentration.
These observations have been interpreted in terms of a
two-stage reaction sequence in which the enzyme (1)
rapidly reacts with the p-nitrophenylacetate to release
NO2
p-Nitrophenolate
p-Nitrophenolate
CH3
C
Enzyme
Acyl–enzyme intermediate
H2O
slow
H+
O
CH3
C
O–
Acetate
+
Enzyme
Enzyme
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Section 15–3. Serine Proteases
Chymotrypsin evidently follows a Ping Pong Bi Bi mechanism (Section 14-5A). Chymotrypsin-catalyzed amide
hydrolysis has been shown to follow a reaction pathway
similar to that of ester hydrolysis but with the first step of
the reaction, enzyme acylation, being rate determining
rather than the deacylation step.
types of nerve cells (Sections 12-4D and 20-5C). The inactivation of acetylcholinesterase prevents the otherwise
rapid hydrolysis of the acetylcholine released by a nerve
impulse and thereby interferes with the regular sequence
of nerve impulses. DIPF is of such great toxicity to humans
that it has been used militarily as a nerve gas. Related compounds, such as parathion and malathion,
b. Identification of the Catalytic Residues
Chymotrypsin’s catalytically important groups were
identified by chemical labeling studies. These are described
below.
O
O 2N
Ser 195. A diagnostic test for the presence of the active
Ser of serine proteases is its reaction with diisopropylphosphofluoridate (DIPF):
+
F
CH3
O
CH2
CH3
O
Diisopropylphosphofluoridate (DIPF)
CH(CH3)2
O
O
P
O
CH2CH3
CH
CH2
C
O
S
CH2
O
CH3
P
S
O
CH3
are useful insecticides because they are far more toxic to
insects than to mammals.
CH(CH3)2
CH2
S
Malathion
O
(Active Ser)
P
C
O
P
CH2CH3
O
O
CH2OH
O
Parathion
CH(CH3)2
(Active Ser)
517
+
O
HF
O
His 57. A second catalytically important residue was discovered through affinity labeling. In this technique, a substrate analog bearing a reactive group specifically binds at
the enzyme’s active site, where it reacts to form a stable
covalent bond with a nearby susceptible group (these
reactive substrate analogs have therefore been described
as the “Trojan horses” of biochemistry). The affinity labeled groups can subsequently be identified by peptide
mapping (Section 7-1K). Chymotrypsin specifically binds
tosyl-L-phenylalanine chloromethyl ketone (TPCK),
CH(CH3)2
DIP–Enzyme
which irreversibly inactivates the enzyme. Other Ser
residues, including those on the same protein, do not react
with DIPF. DIPF reacts only with Ser 195 of chymotrypsin,
thereby demonstrating that this residue is the enzyme’s active
Ser.
The use of DIPF as an enzyme inactivating agent came
about through the discovery that organophosphorus compounds such as DIPF are potent nerve poisons. The neurotoxicity of DIPF arises from its ability to inactivate
acetylcholinesterase, a serine esterase that catalyzes the
hydrolysis of acetylcholine:
O
+
(CH3)3N
CH2
CH2
O
O
CH3
+
CH3
C
H2O
CH2
Choline
OH
+
H
CH2
C
CH3
Acetylcholine is a neurotransmitter: It transmits nerve impulses across the synapses (junctions) between certain
CH2Cl
Cl
HCl
CH2
N
CH2
N
O
–O
C
Chymotrypsin
N
O
CH
O
Chymotrypsin
acetylcholinesterase
CH2
NH
because of its resemblance to a Phe residue (one of chymotrypsin’s preferred residues; Table 7-2). Active site–
bound TPCK’s chloromethyl ketone group is a strong
alkylating agent; it reacts with His 57 (Fig. 15-19), thereby
Acetylcholine
+
(CH3)3N
S
CH2 O
His 57
+
C
R
TPCK
O
N
CH2
C
O
R
FIGURE 15-19
alkylate His 57.
Reaction of TPCK with chymotrypsin to
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Chapter 15. Enzymatic Catalysis
(a)
(a)
X-Ray structure of bovine trypsin. (a) A
drawing of the enzyme in complex with a polypeptide substrate
(green) that has its Arg side chain occupying the enzyme’s
specificity pocket (stippling). The C backbone of the enzyme is
shown together with its disulfide bonds and the side chains of
the catalytic triad, Ser 195, His 57, and Asp 102. The active
sites of chymotrypsin and elastase contain almost identically
arranged catalytic triads. [Illustration, Irving Geis/Geis
Archives Trust. Copyright Howard Hughes Medical Institute.
FIGURE 15-20
Reproduced with permission.] (b) A ribbon diagram of trypsin
highlighting its secondary structure and indicating the
arrangement of its catalytic triad. (c) A drawing showing the
surface of trypsin (blue) superimposed on its polypeptide
backbone ( purple). The side chains of the catalytic triad are
shown in green. [Courtesy of Arthur Olson, The Scripps
Research Institute, La Jolla, California.] Parts a, b, and c have
See Kinemage
approximately the same orientation.
Exercise 10-1
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Section 15–3. Serine Proteases
amino acid residue numbering scheme. Bovine chymotrypsin is synthesized as an inactive 245-residue precursor
named chymotrypsinogen that is proteolytically converted
to chymotrypsin (Section 15-3E). In what follows, the numbering of the amino acid residues in chymotrypsin, trypsin,
and elastase will be that of the corresponding residues in
bovine chymotrypsinogen.
The X-ray structure of bovine chymotrypsin was elucidated in 1967 by David Blow. This was followed by the determination of the structures of bovine trypsin (Fig. 15-20)
by Robert Stroud and Richard Dickerson, and porcine
elastase by David Shotton and Herman Watson. Each of
these proteins is folded into two domains, both of which
have extensive regions of antiparallel -sheets in a barrellike arrangement but contain little helix. The catalytically
essential His 57 and Ser 195 are located at the substratebinding site together with the invariant (in all serine proteases) Asp 102, which is buried in a solvent-inaccessible
pocket. These three residues form a hydrogen bonded constellation referred to as the catalytic triad (Figs. 15-20 and
15-21).
inactivating the enzyme. The TPCK reaction is inhibited
by -phenylpropionate,
CH2
CH2
COO–
-Phenylpropionate
a competitive inhibitor of chymotrypsin that presumably
competes with TPCK for its enzymatic binding site.
Moreover, the TPCK reaction does not occur in 8M urea,
a denaturing reagent, or with DIP–chymotrypsin, in which
the active site is blocked. These observations establish that
His 57 is an essential active site residue of chymotrypsin.
B. X-Ray Structures
Bovine chymotrypsin, bovine trypsin, and porcine elastase
are strikingly homologous: The primary structures of these
240-residue monomeric enzymes are 40% identical and
their internal sequences are even more alike (in comparison, the and chains of human hemoglobin have a 44%
sequence identity). Furthermore, all of these enzymes have
an active Ser and a catalytically essential His as well as similar kinetic mechanisms. It therefore came as no surprise
when their X-ray structures all proved to be closely related.
To most conveniently compare the structures of these
three digestive enzymes, they have been assigned the same
a. The Structural Basis of Substrate Specificity Can Be
Quite Complex
The X-ray structures of the above three enzymes suggest the basis for their differing substrate specificities
(Table 7-2):
1. In chymotrypsin, the bulky aromatic side chain of
the preferred Phe, Trp, or Tyr residue that contributes the
C
His 57
Ser 195
N
Asp 102
(b)
FIGURE 15-20
(c)
(Continued)
519
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Chapter 15. Enzymatic Catalysis
Gly 193
Ser 195
Catalytic
triad
Asp 194
His 57
Ile 16
Asp 102
carbonyl group of the scissile peptide fits snugly into a slitlike hydrophobic pocket, the specificity pocket, that is located near the catalytic groups (Fig. 15-20a).
2. In trypsin, the residue corresponding to chymotrypsin Ser 189, which lies at the back of the specificity
pocket, is the anionic residue Asp. The cationic side chains
of trypsin’s preferred residues, Arg or Lys, can therefore
form ion pairs with this Asp residue. The rest of chymotrypsin’s specificity pocket is preserved in trypsin so that it
can accommodate the bulky side chains of Arg and Lys.
3. Elastase is so named because it rapidly hydrolyzes
the otherwise nearly indigestible Ala, Gly, and Val-rich
protein elastin (a connective tissue protein with rubberlike
elastic properties). Elastase’s specificity pocket is largely
occluded by the side chains of a Val and a Thr residue that
replace two Gly’s lining this pocket in both chymotrypsin
and trypsin. Consequently elastase, whose specificity
pocket is better described as a depression, specifically
cleaves peptide bonds after small neutral residues, particularly Ala. In contrast, chymotrypsin and trypsin hydrolyze
such peptide bonds extremely slowly because these small
substrates cannot be sufficiently immobilized on the enzyme surface for efficient catalysis to occur (Section
15-1E).
Thus, for example, trypsin catalyzes the hydrolysis of peptidyl amide substrates with an Arg or Lys residue preceding
the scissile bond with an efficiency, as measured by kcatKM
(Section 14-2B), that is 106-fold greater than that for
the corresponding Phe-containing substrates. Conversely,
chymotrypsin catalyzes the hydrolysis of substrates after
Phe, Trp, and Tyr residues 104-fold more efficiently than
after the corresponding Lys-containing substrates.
Despite the foregoing, the mutagenic change in trypsin
of Asp 189 S Ser (D189S) by William Rutter did not switch
its specificity to that of chymotrypsin but instead yielded
FIGURE 15-21 The active site residues of chymotrypsin. The
view is in approximately the same direction as in Fig. 15-20.
The catalytic triad consists of Ser 195, His 57, and Asp 102.
[After Blow, D.M. and Steitz, T.A., Annu. Rev. Biochem. 39, 86
(1970).]
a poor, nonspecific protease. Moreover, even replacing the
other three residues in trypsin’s specificity pocket that
differ from those in chymotrypsin, with those of chymotrypsin, fails to yield a significantly improved enzyme.
However, trypsin is converted to a reasonably active
chymotrypsin-like enzyme when, in addition to the foregoing changes (collectively designated S1), both of its
two surface loops that connect the walls of the specificity pocket, L1 (residues 185–188) and L2 (residues
221–225), are replaced by those of chymotrypsin (termed
TrSCh[S1L1L2]). Although this mutant enzyme still
has a low substrate-binding affinity, KS, the additional mutation Y172W in a third surface loop yields an enzyme
(TrSCh[S1L1L2+Y172W]) that has 15% of chymotrypsin’s catalytic efficiency. Curiously, these loops, whose
sequences are largely conserved in each enzyme, are not
structural components of either the specificity pocket or
the extended substrate binding site in chymotrypsin or in
trypsin (Fig. 15-20a).
Careful comparisons, by Charles Craik and Robert
Fletterick, of the X-ray structures of chymotrypsin and
trypsin with those of the closely similar TrSCh[S1L1
L2] and TrSCh[S1L1L2Y172W] in complex with a
Phe-containing chloromethyl ketone inhibitor reveal the
structural basis of substrate specificity in trypsin and
chymotrypsin. Efficient catalysis in the serine proteases requires that the enzyme’s active site be structurally intact
and that the substrate’s scissile bond be properly positioned
relative to the catalytic triad and other components of the
active site (see below). The above mutagenic changes do
not affect the structure of the catalytic triad or those portions of the active site that bind the substrate’s leaving
group (that segment on the C-terminal side of the scissile
bond). However, the main chain conformation of the conserved Gly 216 (which forms two hydrogen bonds to the
backbone of the third residue before the substrate’s scis-
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Section 15–3. Serine Proteases
sile bond in an antiparallel pleated sheet–like arrangement) differs in trypsin and chymotrypsin and adopts a
chymotrypsin-like structure in both hybrid proteins.
Evidently, if Gly 216 adopts a trypsin-like conformation,
the scissile bond in Phe-containing substrates is misoriented for efficient catalysis. Thus, despite the fact that Gly
216 is conserved in trypsin and chymotrypsin, the differing
structures of loop L2 in the two enzymes maintain it in distinct conformations.
Loop L1, which interacts with L2 in both trypsin and
chymotrypsin, is largely disordered in the X-ray structure
of TrSCh[S1L1L2]. Modeling a trypsin-like L1 into
TrSCh[S1L1L2] results in severe steric clashes with
the chymotrypsin-like L2. Thus, the requirement of a chymotrypsin-like L1 for the efficient catalysis by TrS
Ch[S1L1L2] appears to arise from the need to permit
L2 to adopt a chymotrypsin-like conformation.
Residue 172 is located at the base of the specificity
pocket. The improvement in substrate binding affinity of
TrSCh[S1L1L2Y172W] over TrSCh[S1L1L2]
arises from structural rearrangements in this region of the
enzyme caused by the increased bulk and different
hydrogen bonding requirements of Trp versus Tyr. These
changes appear to improve both the structural stability of
residues forming the specificity pocket and their specificity
for chymotrypsin-like substrates. These results therefore
highlight an important caveat for genetic engineers:
Enzymes are so exquisitely tailored to their functions that
they often respond to mutagenic tinkering in unexpected
ways.
b. Evolutionary Relationships among Serine Proteases
We have seen that sequence and structural homologies
among proteins reveal their evolutionary relationships
(Sections 7-3 and 9-6). The great similarities among chymotrypsin, trypsin, and elastase indicate that these proteins
evolved through gene duplications of an ancestral serine
protease followed by the divergent evolution of the resulting enzymes (Section 7-3C).
Several serine proteases from various sources provide
further insights into the evolutionary relationships among
the serine proteases. Streptomyces griseus protease A
(SGPA) is a bacterial serine protease of chymotryptic
specificity that exhibits extensive structural similarity, although only 20% sequence identity, with the pancreatic
serine proteases. The primordial trypsin gene evidently
arose before the divergence of prokaryotes and eukaryotes.
There are three known serine proteases whose primary
and tertiary structures bear no discernible relationship to
each other or to chymotrypsin but which, nevertheless,
contain catalytic triads at their active sites whose structures
closely resemble that of chymotrypsin:
Subtilisin
NH+3
521
ClpP
Serine
Chymotrypsin carboxypeptidase II protease
NH+3
NH+3
NH+3
Asp 32
His 64
His 57
Ser
146
Asp
102
Ser 125
Leu 126
Gly 127
Ser
97
Asp
338
Ser 195
Ser 221
Ser 214
Trp 215
Gly 216
COO–
COO–
His
397
COO–
His
122
Asp
171
COO–
Relative positions of the active site residues in
subtilisin, chymotrypsin, serine carboxypeptidase II, and ClpP
protease. The peptide backbones of Ser 214, Trp 215, and Gly
216 in chymotrypsin, and their counterparts in subtilisin,
participate in substrate-binding interactions. [After Robertus,
J.D., Alden, R.A., Birktoft, J.J., Kraut, J., Powers, J.C., and
See Kinemage
Wilcox, P.E., Biochemistry 11, 2449 (1972).]
FIGURE 15-22
Exercise 10-2
carboxypeptidase A (Fig. 8-19a) even though the latter
protease has an entirely different catalytic mechanism from
that of the serine proteases (see Problem 3).
3. E. coli ClpP, which functions in the degradation of
cellular proteins (Section 32-6B).
Since the orders of the corresponding active site residues
in the amino acid sequences of the four types of serine
proteases are quite different (Fig. 15-22), it seems highly
improbable that they could have evolved from a common
ancestor serine protease. These proteins apparently constitute a remarkable example of convergent evolution: Nature
seems to have independently discovered the same catalytic
mechanism at least four times. (In addition, human cytomegalovirus protease, an essential protein for virus replication that bears no resemblance to the above proteases,
has active site Ser and His residues whose relative positions are similar to those in other serine proteases but lacks
an active site Asp residue; it appears to have a catalytic
dyad.)
C. Catalytic Mechanism
1. Subtilisin, an endopeptidase that was originally isolated from Bacillus subtilis.
See Guided Exploration 12: The Catalytic Mechanism of Serine
Proteases The extensive active site homologies among the
2. Wheat germ serine carboxypeptidase II, an exopeptidase whose structure is surprisingly similar to that of
various serine proteases indicate that they all have the
same catalytic mechanism. On the basis of considerable
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Chapter 15. Enzymatic Catalysis
522
1. After chymotrypsin has bound substrate to form the
Michaelis complex, Ser 195, in the reaction’s rate-determining step, nucleophilically attacks the scissile peptide’s
carbonyl group to form a complex known as the tetrahedral
chemical and structural data gathered in many laboratories, the following catalytic mechanism has been formulated for the serine proteases, here given in terms of chymotrypsin (Fig. 15-23):
His
57
Asp
102
H 2C
CH2
H
....
N1
O
..
CH2
H
O
3
N+
1
C
O–
H
Tetrahedral intermediate
2
His
57
Asp
102
His
57
CH2
H 2C
Ser
195
N1
CH2
.O
...
H
C –
....
....
H
O
R
N
Enzyme–substrate
complex
.O
H2C
...
C –
O
R
O
H
CH2
H
Nucleophilic
attack
C
N
Asp
102
Ser
195
N1
R
R
Substrate
polypeptide
H
O
3
N
CH2
.O
–
C
...
H 2C
Ser
195
....
.O
C –
...
His
57
Asp
102
H2O
Ser
195
N1
O
3
2
3
CH2
N
CH2
N
O
H
R
H
R
R
C
N
C
O
O
H
New N-terminus of
cleaved polypeptide
chain
RNH2
H
O
O
Acyl–enzyme intermediate
3
.
....
C –
O
CH2
H 2C
H
Ser
195
N
N+
Ser
195
H
N
CH2
N
CH2
H
O
H
R
O
C
H
O–
Tetrahedral intermediate
Catalytic mechanism of the serine proteases.
The reaction involves (1) the nucleophilic attack of the active
site Ser on the carbonyl carbon atom of the scissile peptide
bond to form the tetrahedral intermediate; (2) the
decomposition of the tetrahedral intermediate to the acyl–
enzyme intermediate through general acid catalysis by the
FIGURE 15-23
C –
O
4
CH2
.O
...
....
H2C
O
...
His
57
Asp
102
His
57
Asp
102
O
+
New C-terminus
of cleaved polypeptide
chain
O
H
R
C
O
Active enzyme
active site Asp-polarized His, followed by loss of the amine
product and its replacement by a water molecule; (3) the
reversal of Step 2 to form a second tetrahedral intermediate; and
(4) the reversal of Step 1 to yield the reaction’s carboxyl
product and the active enzyme.
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Section 15–3. Serine Proteases
intermediate (covalent catalysis). X-Ray studies indicate
that Ser 195 is ideally positioned to carry out this nucleophilic attack (proximity and orientation effects). The imidazole ring of His 57 takes up the liberated proton, thereby
forming an imidazolium ion (general base catalysis). This
process is aided by the polarizing effect of the unsolvated
carboxylate ion of Asp 102, which is hydrogen bonded to
His 57 (electrostatic catalysis; see Section 15-3D). Indeed,
the mutagenic replacement of trypsin’s Asp 102 by Asn
leaves the enzyme’s KM substantially unchanged at neutral
pH but reduces its kcat to 0.05% of its wild-type value.
Neutron diffraction studies have demonstrated that Asp
102 remains a carboxylate ion rather than abstracting a proton from the imidazolium ion to form an uncharged carboxylic acid group. The tetrahedral intermediate has a
well-defined, although transient, existence. We shall see
that much of chymotrypsin’s catalytic power derives from
its preferential binding of the transition state leading to this
intermediate (transition state binding catalysis).
523
The portion of BPTI in contact with the trypsin active
site resembles bound substrate. The side chain of BPTI Lys
15I (here “I” differentiates BPTI residues from trypsin
residues) occupies the trypsin specificity pocket (Fig.
15-24a) and the peptide bond between Lys 15I and Ala 16I
is positioned as if it were the scissile peptide bond
(Fig. 15-24b). What is most remarkable about this structure
is that its active site complex assumes a conformation well
along the reaction coordinate toward the tetrahedral
intermediate: The side chain oxygen of trypsin Ser 195, the
active Ser, is in closer-than-van der Waals contact (2.6 Å)
with the pyramidally distorted carbonyl carbon of BPTI’s
“scissile” peptide. Despite this close contact, the proteolytic
reaction cannot proceed past this point along the reaction
coordinate because of the rigidity of the active site complex and because it is so tightly sealed that the leaving group
cannot leave and water cannot enter the reaction site.
Protease inhibitors are common in nature, where they have
protective and regulatory functions. For example, certain
2. The tetrahedral intermediate decomposes to the
acyl–enzyme intermediate under the driving force of proton donation from N3 of His 57 (general acid catalysis).
The amine leaving group (RNH2, the new N-terminal portion of the cleaved polypeptide chain) is released from the
enzyme and replaced by water from the solvent.
3 & 4. The acyl-enzyme intermediate (which, in the absence of enzyme, would be a stable compound) is rapidly
deacylated by what is essentially the reverse of the previous steps followed by the release of the resulting carboxylate product (the new C-terminal portion of the cleaved
polypeptide chain), thereby regenerating the active enzyme. In this process, water is the attacking nucleophile
and Ser 195 is the leaving group.
D. Testing the Catalytic Mechanism
The formulation of the foregoing model for catalysis by
serine proteases has prompted numerous investigations of
its validity. In this section we discuss several of the most
revealing of these studies.
(a)
Ser 195
O
H
Ala 16I
O
a. The Tetrahedral Intermediate Is Mimicked in a
Complex of Trypsin with Trypsin Inhibitor
Convincing structural evidence for the existence of the
tetrahedral intermediate was provided by Robert Huber
in an X-ray study of the complex between bovine pancreatic trypsin inhibitor (BPTI) and trypsin. The 58-residue
protein BPTI, whose folding pathway we examined in
Section 9-1C, binds to and inactivates trypsin, thereby preventing any trypsin that is prematurely activated in the
pancreas from digesting that organ (see Section 15-3E).
BPTI binds to the active site region of trypsin across
a tightly packed interface that is cross-linked by a complex network of hydrogen bonds. This complex’s 1013M1
association constant, among the largest of any known protein–protein interaction, emphasizes BPTI’s physiological
importance.
C
C
C
(b)
(b)
N
H
Lys 15I
FIGURE 15-24 Trypsin–BPTI complex. (a) The X-ray
structure shown as a cutaway surface drawing indicating how
trypsin (red) binds BPTI (green). The green protrusion
extending into the red cavity near the center of the figure
represents the Lys 15I side chain occupying trypsin’s specificity
pocket. Note the close complementary fit of these two proteins.
[Courtesy of Michael Connolly, New York University.]
(b) Trypsin Ser 195, the active Ser, is in closer-than-van der
Waals contact with the carbonyl carbon of BPTI’s scissile
peptide, which is pyramidally distorted toward Ser 195. The
normal proteolytic reaction is apparently arrested somewhere
along the reaction coordinate between the Michaelis complex
and the tetrahedral intermediate.
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Chapter 15. Enzymatic Catalysis
plants release protease inhibitors in response to insect
bites, thereby causing the offending insect to starve by inactivating its digestive enzymes. Protease inhibitors constitute 10% of the nearly 200 proteins of blood serum. For
instance, 1-proteinase inhibitor, which is secreted by the
liver, inhibits leukocyte elastase (leukocytes are a type of
white blood cell; the action of leukocyte elastase is thought
to be part of the inflammatory process). Pathological variants of 1-proteinase inhibitor with reduced activity are associated with pulmonary emphysema, a degenerative disease of the lungs resulting from the hydrolysis of its elastic
fibers. Smokers also suffer from reduced activity of their
1-proteinase inhibitor because of the oxidation of its active site Met residue. Full activity of this inhibitor is not regained until several hours after smoking.
b. Serine Proteases Preferentially Bind the
Transition State
Detailed comparisons of the X-ray structures of several
serine protease–inhibitor complexes have revealed a further structural basis for catalysis in these enzymes (Fig.
15-25):
1. The conformational distortion that occurs with the
formation of the tetrahedral intermediate causes the carbonyl oxygen of the scissile peptide to move deeper into
the active site so as to occupy a previously unoccupied position, the oxyanion hole.
2. There it forms two hydrogen bonds with the enzyme
that cannot form when the carbonyl group is in its normal
trigonal conformation. These two enzymatic hydrogen
bond donors were first noted by Joseph Kraut to occupy
corresponding positions in chymotrypsin and subtilisin. He
proposed the existence of the oxyanion hole based on the
premise that convergent evolution had made the active
sites of these unrelated enzymes functionally identical.
3. The tetrahedral distortion, moreover, permits the
formation of an otherwise unsatisfied hydrogen bond between the enzyme and the backbone NH group of the
residue preceding the scissile peptide. Consequently, the
enzyme binds the tetrahedral intermediate in preference to
either the Michaelis complex or the acyl–enzyme intermediate.
It is this phenomenon that is responsible for much of the
catalytic efficiency of serine proteases (see below). In fact,
the reason that DIPF is such an effective inhibitor of serine proteases is because its tetrahedral phosphate group
makes this compound a transition state analog of the
enzyme.
c. The Tetrahedral Intermediate and the Water
Molecule Attacking the Acyl–Enzyme Intermediate
Have Been Directly Observed
Most enzymatic reactions turn over far too rapidly for
their intermediate states to be studied by X-ray or NMR
techniques. Consequently, much of our structural knowledge of these intermediate states derives from the study of
enzyme–inhibitor complexes or complexes of substrates
with inactivated enzymes. Yet the structural relevance of
these complexes is subject to doubt precisely because they
are catalytically unproductive.
In an effort to rectify this situation for serine proteases,
Janos Hadju and Christopher Schofield searched for
peptide–protease complexes that are stable at pH’s at
which the protease is inactive but which could be rendered
active by changing the pH. To do so, they screened libraries
(b)
(a)
Oxyanion hole
Ser 195
Ser 195
His 57
N
H
O
Cα
Cβ
.
C
R
...
O
H
O
NH
Gly 193
....
Gly 193
HN
N
O
NH ...–O
C
Asp
102
R′
NH
... O–
R
N
H
O
C
Gly 193
Gly 193
Transition state stabilization in the serine
proteases. (a) In the Michaelis complex, the trigonal carbonyl
carbon of the scissile peptide is conformationally constrained
from binding in the oxyanion hole (upper left). (b) In the
tetrahedral intermediate, the now charged carbonyl oxygen of
the scissile peptide (the oxyanion) has entered the oxyanion
hole, thereby hydrogen bonding to the backbone NH groups of
O
O
C
C
FIGURE 15-25
His 57
N
H
...
N
H
Cα
HN ... H
Cβ
–O
N + N..H..
C
Asp
102
R′
Gly 193 and Ser 195. The consequent conformational distortion
permits the NH group of the residue preceding the scissile
peptide bond to form an otherwise unsatisfied hydrogen bond
to Gly 193. Serine proteases therefore preferentially bind the
tetrahedral intermediate. [After Robertus, J.D., Kraut, J.,
Alden, R.A., and Birktoft, J.J., Biochemistry 11, 4302 (1972).]
See Kinemage Exercise 10-3
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Section 15–3. Serine Proteases
525
of peptides for their ability to bind to porcine pancreatic
elastase at pH 3.5 (at which pH His 57 is protonated and
hence unable to act as a general base) through the use of
ESI-MS (Section 7-1J). They thereby discovered that
YPFVEPI, a heptapeptide segment of the human milk protein -casein that is named BCM7, forms a complex with
elastase, whose mass is consistent with the formation of an
ester linkage between BCM7 and the enzyme. In the presence of 18OH2 at pH 7.5 (where elastase is active), the 18O
label was incorporated into both BCM7 and the elastase–
BCM7 complex, thereby demonstrating that the reaction
of BCM7 with elastase is reversible at this pH.
Fragmentation studies by fast atom bombardment–tandem
mass spectroscopy (FAB–MS/MS; Section 7-1J) further revealed that BCM7 that had been incubated with elastase
in the presence of 18OH2 at pH 7.5 incorporated the 18O
label into only its C-terminal Ile residue.
The X-ray structure of the BCM7–elastase complex at
pH 5 (Fig. 15-26a) revealed that BCM7’s C-terminal carboxyl group, in fact, forms an ester linkage with elastase’s
Ser 195 side chain hydroxyl group to form the expected
acyl–enzyme intermediate. Moreover, this X-ray structure
reveals the presence of a bound water molecule that appears poised to nucleophilically attack the ester linkage
(the distance from this water molecule to BCM7’s Cterminal C atom is 3.1 Å and the line between them is
nearly perpendicular to the plane of the acyl group). His
57, which is hydrogen bonded to this water molecule, is
properly positioned to abstract one of its protons, thereby
activating it for the nucleophilic attack (general base catalysis). The carbonyl O atom of the acyl group occupies
the enzyme’s oxyanion hole such that it is hydrogen bonded
to the main chain N atoms of both Ser 195 and Gly 193.
This is in agreement with spectroscopic measurements indicating that the acyl–enzyme intermediate’s carbonyl
group is, in fact, hydrogen bonded to the oxyanion hole.
It was initially assumed that the oxyanion hole acts only
to stabilize the tetrahedral oxyanion transition state that
resides near the tetrahedral intermediate on the catalytic
reaction coordinate. However, it now appears that the
oxyanion hole also functions to polarize the carbonyl group
of the acyl–enzyme intermediate toward an oxyanion
(electrostatic catalysis).
The catalytic reaction was initiated in crystals of the
BCM7–elastase complex by transferring them to a buffer
at pH 9. After soaking in this buffer for 1 min, the crystals
were rapidly frozen in liquid N2 (196°C), thereby arresting the enzymatic reaction (recall that the catalytically
essential collective motions of proteins cease at such low
temperatures; Section 9-4). The X-ray structure of such a
frozen crystal (Fig. 15-26b) revealed that the above acyl–
enzyme intermediate had converted to the tetrahedral in-
(a)
(b)
X-Ray structures of porcine pancreatic elastase
in complex with the heptapeptide BCM7 (YPFVEPI). The
residues of elastase are specified by the three-letter code and
those of BCM7 are specified by the one-letter code. (a) The
complex at pH 5. The enzyme’s active site residues and the
heptapeptide (whose N-terminal three residues are disordered)
are shown in ball-and-stick form with elastase C green, BCM7
C cyan, N blue, O red, S yellow, and the bond between the Ser
195 O atom and the C-terminal C atom of BCM7 magenta. The
enzyme-bound water molecule, which appears poised to
nucleophilically attack the acyl–enzyme’s carbonyl C atom, is
represented by an orange sphere. The dashed gray lines
represent the catalytically important hydrogen bonds and the
dotted gray line indicates the trajectory that the bound water
molecule presumably follows in nucleophilically attacking the
acyl group’s carbonyl C atom. (b) The complex after being
brought to pH 9 for 1 min and then rapidly frozen in liquid
nitrogen. The various groups in the structure are represented
and colored as in Part a. Note that the water molecule in
Part a has become a hydroxyl substituent (orange) to the
carbonyl C atom, thereby yielding the tetrahedral intermediate.
[Based on X-ray structures by Christopher Schofield and Janos
Hadju, University of Oxford, U.K. PDBids (a) 1HAX and (b)
1HAZ.]
FIGURE 15-26
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Chapter 15. Enzymatic Catalysis
termediate, whose oxyanion, as expected, remained hydrogen bonded to the N atoms of Ser 195 and Gly 193.
Comparison of this crystal structure with that of the acyl–
enzyme intermediate reveals that the enzyme’s active site
residues do not significantly change their positions in the
conversion from the acyl–enzyme intermediate to the
tetrahedral intermediate. However, the peptide substrate
must do so out of steric necessity when the trigonal planar
acyl group converts to the tetrahedral oxyanion (compare
Figs. 15-26a and 15-26b). In response, several enzyme
residues that contact the peptide but which are distant from
the active site also shift their positions (not shown in Fig.
15-26).
d. The Role of the Catalytic Triad: Low-Barrier
Hydrogen Bonds
The earlier literature postulated that the Asp 102polarized His 57 side chain directly abstracts a proton
from Ser 195, thereby converting its weakly nucleophilic
¬ CH2OH group to a highly nucleophilic alkoxide ion,
¬ CH2O:
His
57
Asp
102
H2C
CH2
O
C –
H
O
Ser
195
N
CH2
N
H
His
57
Asp
102
CH2
O
H2C
O
C
O
H
Ser
195
N
CH2
N
H
–
O
"Charge relay system"
In the process, the anionic charge of Asp 102 was thought
to be transferred, via a tautomeric shift of His 57, to Ser
195. The catalytic triad was therefore originally named the
charge relay system. It is now realized, however, that such
a mechanism is implausible because an alkoxide ion
(pK 15) has far greater proton affinity than does His 57
(pK L 7, as measured by NMR techniques). How, then,
can Asp 102 nucleophilically activate Ser 195?
A possible solution to this conundrum has been pointed
out by W.W. Cleland and Maurice Kreevoy and, independently, by John Gerlt and Paul Gassman. Proton transfers
between hydrogen bonded groups (D¬ H p A) only occur at physiologically reasonable rates when the pK of the
proton donor is no more than 2 or 3 pH units greater than
that of the protonated form of the proton acceptor (the
height of the kinetic barrier, G ‡, for the protonation of
an acceptor by a more basic donor increases with the dif-
ference between the pK’s of the donor and acceptor).
However, when the pK’s of the hydrogen bonding donor
(D) and acceptor (A) groups are nearly equal, the distinction between them breaks down: The hydrogen atom
becomes more or less equally shared between them
(D p H p A). Such low-barrier hydrogen bonds (LBHBs)
are unusually short and strong (they are also known as
short, strong hydrogen bonds): They have, as studies of
model compounds in the gas phase indicate, association
free energies as high as 40 to 80 kJ mol1 versus the
12 to 30 kJ mol1 for normal hydrogen bonds (the energy of the normally covalent D¬ H bond is subsumed
into the low-barrier hydrogen bonding system) and a
D p A length of 2.55 Å for O¬ H p O and 2.65 Å for
N¬ H p O versus 2.8 to 3.1 Å for normal hydrogen bonds.
LBHBs are unlikely to exist in dilute aqueous solution
because water molecules, which are excellent hydrogen
bonding donors and acceptors, effectively compete with
D¬ H and A for hydrogen bonding sites. However,
LBHBs may exist in nonaqueous solution and in the active
sites of enzymes that exclude bulk solvent water. If so, an
effective enzymatic “strategy” would be to convert a weak
hydrogen bond in the Michaelis complex to a strong hydrogen bond in the transition state, thereby facilitating proton transfer while applying the difference in the free energy between the normal and low-barrier hydrogen bonds
to preferentially binding the transition state. In fact, as
Perry Frey has shown, the NMR spectrum of the proton
linking His 57 to Asp 102 in chymotrypsin (which exhibits
a particularly large downfield chemical shift indicative of
deshielding) is consistent with the formation of an LBHB
in the transition state (see Fig. 15-25b; the pK’s of protonated His 57 and Asp 102 are nearly equal in the anhydrous environment of the active site complex). This presumably promotes proton transfer from Ser 195 to His 57
as in the charge relay mechanism. Moreover, an ultrahigh
(0.78 Å) resolution X-ray structure of Bacillus lentus
subtilisin by Richard Bott reveals that the hydrogen bond
between His 64 and Asp 32 of its catalytic triad has an unusually short N p O distance of 2.62 0.01 Å and that its
H atom is nearly centered between the N and O atoms
(note that this highly accurate protein X-ray structure is
one of the very few in which H atoms are observed and in
which short D p A distances are confidently measured).
Although several studies, such as the foregoing, have
revealed the existence of unusually short hydrogen bonds
in enzyme active sites, it is far more difficult to demonstrate experimentally that they are unusually strong, as
LBHBs are predicted to be. In fact, several studies of the
strengths of unusually short hydrogen bonds in organic
model compounds in nonaqueous solutions suggest that
these hydrogen bonds are not unusually strong. Consequently, a lively debate has ensued as to the catalytic significance of LBHBs. Yet if enzymes do not form LBHBs,
it remains to be explained how, in numerous widely accepted enzymatic mechanisms that we shall encounter, the
conjugate base of an acidic group can abstract a proton
from a far more basic group.
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Section 15–3. Serine Proteases
e. Much of a Serine Protease’s Catalytic Activity Arises
from Preferential Transition State Binding
Despite the foregoing, blocking the action of the catalytic triad through the specific methylation of His 57 by
treating chymotrypsin with methyl-p-nitrobenzene sulfonate
His
57
CH2
O
H
N1
+
3
O2N
N
S
O
CH3
O
Methyl-p-nitrobenzene
sulfonate
His
57
CH2
H
N1
+
3
O2N
SO3–
N+
CH3
yields an enzyme that is a reasonably good catalyst: It enhances the rate of proteolysis by as much as a factor of
2 106 over the uncatalyzed reaction, whereas the native
enzyme has a rate enhancement factor of 1010. Similarly,
the mutation of Ser 195, His 57, or even all three residues
of the catalytic triad yields enzymes that enhance proteolysis rates by 5 104-fold over that of the uncatalyzed reaction. Evidently, the catalytic triad provides a nucleophile
and is an alternate source and sink of protons (general acid–
base catalysis). However, a large portion of chymotrypsin’s
rate enhancement must be attributed to its preferential binding of the catalyzed reaction’s transition state.
E. Zymogens
Most proteolytic enzymes are biosynthesized as somewhat
larger inactive precursors known as zymogens (enzyme
precursors, in general, are known as proenzymes). In the
case of digestive enzymes, the reason for this is clear: If
these enzymes were synthesized in their active forms, they
would digest the tissues that synthesized them. Indeed,
acute pancreatitis, a painful and sometimes fatal condition
that can be precipitated by pancreatic trauma, is characterized by the premature activation of the digestive enzymes synthesized by this gland.
a. Serine Proteases Are Autocatalytically Activated
Trypsin, chymotrypsin, and elastase are activated according to the following pathways:
Trypsin. The activation of trypsinogen, the zymogen of
trypsin, occurs as a two-stage process when trypsinogen enters the duodenum from the pancreas. Enteropeptidase, a
single-pass transmembrane serine protease that is located
527
in the duodenal mucosa, specifically hydrolyzes trypsinogen’s Lys 15 ¬ Ile 16 peptide bond, thereby excising its Nterminal hexapeptide (Fig. 15-27). This yields the active
enzyme, which has Ile 16 at its N-terminus. Since this activating cleavage occurs at a trypsin-sensitive site (recall
that trypsin cleaves after Arg and Lys residues), the small
amount of trypsin produced by enteropeptidase also catalyzes activation, generating more trypsin, etc.; that is,
trypsinogen activation is autocatalytic.
Chymotrypsin. Chymotrypsinogen is activated by the specific tryptic cleavage of its Arg 15 ¬ Ile 16 peptide bond
to form -chymotrypsin (Fig. 15-28). -Chymotrypsin
subsequently undergoes autolysis (self-digestion) to specifically excise two dipeptides, Ser 14–Arg 15 and Thr 147–
Asn 148, thereby yielding the equally active enzyme -chymotrypsin (heretofore and hereafter referred to as
chymotrypsin). The biochemical significance of this latter
process, if any, is unknown.
Elastase. Proelastase, the zymogen of elastase, is activated
similarly to trypsinogen by a single tryptic cleavage that
excises a short N-terminal polypeptide.
b. Biochemical “Strategies” That Prevent Premature
Zymogen Activation
Trypsin activates pancreatic procarboxypeptidases A
and B and prophospholipase A2 (the action of phospholipase A2 is outlined in Section 25-1) as well as the
pancreatic serine proteases. Premature trypsin activation
can consequently trigger a series of events that lead to
pancreatic self-digestion. Nature has therefore evolved an
elaborate defense against such inappropriate trypsin activation. We have already seen (Section 15-3D) that pancreatic trypsin inhibitor binds essentially irreversibly to any
trypsin formed in the pancreas so as to inactivate it.
Furthermore, the trypsin-catalyzed activation of trypsinogen (Fig. 15-27) occurs quite slowly, presumably because
the unusually large negative charge of its highly evolutionarily conserved N-terminal hexapeptide repels the Asp
at the back of trypsin’s specificity pocket. Finally, pancreatic zymogens are stored in intracellular vesicles called
+
H3N
10
Val
(Asp)4
15
16
Lys
Ile
Val
...
Trypsinogen
enteropeptidase or
trypsin
+
H3N
Val
(Asp)4
Lys
+
Ile
Val
...
Trypsin
Activation of trypsinogen to form trypsin.
Proteolytic excision of the N-terminal hexapeptide is catalyzed
by either enteropeptidase or trypsin. The chymotrypsinogen
residue numbering is used here; that is, Val 10 is actually
trypsinogen’s N-terminus and Ile 16 is trypsin’s N-terminus.
FIGURE 15-27
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Chapter 15. Enzymatic Catalysis
Activation of
chymotrypsinogen by proteolytic cleavage.
Both - and -chymotrypsin are
See Kinemage
enzymatically active.
FIGURE 15-28
Chymotrypsinogen
(inactive)
1
122
Cys
Exercise 10-4
S
Cys
S
136
201
S
Cys
S
245
Cys
trypsin
Arg
π-Chymotrypsin
(active)
1
15
Ile
16
122
136
201
S
S
245
S
S
chymotrypsin
Ser Arg
14
Tyr
Leu
α-Chymotrypsin
(active)
1
c. Zymogens Have Distorted Active Sites
Since the zymogens of trypsin, chymotrypsin, and elastase have all their catalytic residues, why aren’t they enzymatically active? Comparisons of the X-ray structures of
trypsinogen with that of trypsin and of chymotrypsinogen
with that of chymotrypsin show that on activation, the
newly liberated N-terminal Ile 16 residue moves from the
surface of the protein to an internal position, where its free
cationic amino group forms an ion pair with the invariant
anionic Asp 194 (Fig. 15-21). Aside from this change, however, the structures of these zymogens closely resemble
those of their corresponding active enzymes. Surprisingly,
this resemblance includes their catalytic triads, an observation which led to the discovery that these zymogens are
actually enzymatically active, albeit at a very low level.
Careful comparisons of the corresponding enzyme and zymogen structures, however, revealed the reason for this
low activity: The zymogens’ specificity pockets and oxyanion holes are improperly formed such that, for example,
the amide NH of chymotrypsin’s Gly 193 points in the wrong
direction to form a hydrogen bond with the tetrahedral intermediate (see Fig. 15-25). Hence, the zymogens’ very low
enzymatic activity arises from their reduced ability to bind
substrate productively and to stabilize the tetrahedral intermediate. These observations provide further structural
evidence favoring the role of preferred transition state
binding in the catalytic mechanism of serine proteases.
4 DRUG DESIGN
The improvements in medical care over the past several
decades are, in large measure, attributable to the development of a huge variety of drugs, which have eliminated
+
147
148
Ala
Ile
13
16
122
136
146
149
S
S
zymogen granules whose membranous walls are thought
to be resistant to enzymatic degradation.
15
Thr Asn
201
245
S
S
or greatly relieved numerous human ailments. Such medications include antibiotics (which have enormously reduced the impact of infectious diseases), anti-inflammatory
agents (which reduce the effects of inflammatory diseases
such as arthritis), analgesics and anesthetics (which make
modern surgical techniques possible), agents that reduce
the incidence and severity of cardiovascular disease and
stroke, antidepressants, antipsychotics, agents that inhibit
stomach acid secretion (which prevent stomach ulcers and
heartburn), agents to combat allergies and asthma, immunosuppressants (which make organ transplants possible), agents used for cancer chemotherapy, and a great
variety of other substances.
Early human cultures almost certainly recognized both
the beneficial and toxic effects of indigenous plant and animal products and used many of them as “medications.”
Unfortunately, most of these substances were useless or
even harmful. Although there were sporadic attempts over
the 2500 years preceding the modern era to formulate rational systems of drug discovery, they had little success because they were based mainly on unfounded theories and
superstition (e.g., the doctrine of signatures stated that if
a plant resembles a particular body part, it must be designed by nature to influence that body part) rather than
observation and experiment. Consequently, at the beginning of the 20th century, only three known drugs, apart
from folk medicines, were effective in treating specific diseases: (1) Digitalis, a heart stimulant extracted from the
foxglove plant (Section 20-3A), was used to treat various
heart conditions; (2) quinine (Section 26-4A), obtained
from the bark and roots of the Cinchona tree, was used to
treat malaria; and (3) mercury was used to treat syphilis
(a cure that was often worse than the disease). It was not
until several decades later that the rise of the scientific
method coupled to the rapidly increasing knowledge of
physiology, biochemistry, and chemistry led to effective
methods of drug discovery. In fact, the vast majority of
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Section 15–4. Drug Design
drugs in use today were discovered and developed in the
past three decades.
In this section we discuss the elements of drug discovery and pharmacology (the science of drugs, including their
composition, uses, and effects). The section ends with a
consideration of one of the major successes of modern drug
discovery methods, HIV protease inhibitors.
A. Techniques of Drug Discovery
Most drugs act by modifying the function of a particular
receptor in the body or in an invading pathogen. In most
cases, the receptor is a protein to which the drug specifically binds. It may be an enzyme, a transmembrane channel that transports a specific substance into or out of a cell
(Chapter 20), and/or a protein that participates in an interor intracellular signaling pathway (Chapter 19). In all of
these cases, a substance that in binding to a receptor modulates its function is known as an agonist, whereas a substance that binds to a receptor without affecting its function but blocks the binding of agonists is called an
antagonist. The biochemical and physiological effects of a
drug and its mechanism of action are referred to as its
pharmacodynamics.
a. Drug Discovery Is a Complex Procedure
How are new drugs discovered? Nearly all drugs that
have been in use for over a decade were discovered by
screening large numbers of synthetic compounds and natural products for the desired effect. Drug candidates that
are natural products are usually discovered by the fractionation of the organisms in which they occur, which are
often plants used in folk remedies of the conditions of interest. Humans having the condition whose treatment is
being sought cannot be used as “guinea pigs” in this initial screening process, and even guinea pigs or other laboratory animals such as mice or dogs (if they can be made
to be suitable models of the condition under consideration) are too expensive to use on the many thousands of
compounds that are usually tested. Thus, in vitro screens
are initially used, such as the degree of binding of a drug
candidate to an enzyme that is implicated in a disease of
interest, toxicity toward the target bacteria in the search
for a new antibiotic, or effects on a line of cultured mammalian cells. However, as the number of drug candidates
is winnowed down, more sensitive screens such as testing
in animals are employed.
A drug candidate that exhibits a desired effect is called
a lead compound (or, colloquially, a lead). A good lead
compound binds to its target receptor with a dissociation
constant, KD 1 M. Such a high affinity is necessary to
minimize a drug’s less specific binding to other macromolecules in the body and to ensure that only low doses of the
drug need be taken. For enzyme inhibitors, the dissociation constant is the inhibitor’s KI or K¿I (Section 14-3).
Other common measures of the effect of a drug are the
IC50, the inhibitor concentration at which an enzyme exhibits 50% of its maximal activity; the ED50, the effective
dose of a drug required to produce a therapeutic effect in
50% of a test sample; the TD50, the mean toxic dose
529
required to produce a particular toxic effect in animals;
and the LD50, the mean lethal dose required to kill 50%
of a test sample.
For an inhibitor of an enzyme that follows MichaelisMenten kinetics, the IC50 is determined by measuring the
ratio I /o for several values of [I] at constant [S], where
I is the initial velocity of the enzyme when the inhibitor concentration is [I]. By dividing equation
[14.24] by equation [14.38] with defined according
to equation [14.37], we see that
KM 3 S4
vI
v0
KM 3S4
KM 3 S4
3I4
KM a1 b 3S4
K1
When I /o 0.5 (50% inhibition),
3I4 3 IC50 4 KI a1 3S 4
KM
b
[15.13]
[15.14]
Consequently, if the measurements of I /o are made with
[S] KM, then [IC50] KI.
The ratio TD50ED50 is defined as a drug’s therapeutic
index, the ratio of the dose of the drug that produces toxicity to that which produces the desired effect. It is, of
course, preferable that a drug have a high therapeutic index, but this is not always possible.
b. Cathepsin K Is a Drug Target for Osteoporosis
The development of genomic sequencing techniques
(Section 7-2B) and hence the characterization of tens of
thousands of previously unknown genes is providing an
enormous number of potential drug targets. For example,
osteoporosis, a condition that afflicts postmenopausal
women and elderly men, is characterized by the progressive loss of bone mass leading to a greatly increased frequency of bone fracture, particularly of the hip, spine, and
wrist. Bones consist of a protein matrix that is 90% type
I collagen (Section 8-2B), in which spindle- or plate-shaped
crystals of hydroxyapatite, Ca5(PO4)3OH, are embedded.
Bones are by no means static structures. They undergo
continuous remodeling through the countervailing action
of two types of bone cells: osteoblasts, which synthesize
bone’s organic matrix in which its mineral component is
laid down; and osteoclasts, which solubilize mineralized
bone matrix through the secretion of proteolytic enzymes
into an extracellular bone resorption pit, which is maintained at pH 4.5. The acidic solution dissolves the bone’s
mineral component, thereby exposing its protein matrix to
proteolytic degradation. Osteoporosis arises when bone resorption outstrips bone formation.
In the search for a drug target for osteoporosis, a cDNA
library (Sections 5-5E and 5-5F) was prepared from an
osteoclastoma (a cancer derived from osteoclasts; normally
osteoclasts are very rare cells). Around 4% of these cDNAs
encode a heretofore unknown protease, which was named
cathepsin K (cathepsins are proteases that occur in the lysosome). Further studies, both at the cDNA and protein levels, indicated that cathepsin K is only expressed at high levels in osteoclasts. Microscopic examination of osteoclasts
that had been stained with antibodies directed against
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Chapter 15. Enzymatic Catalysis
cathepsin K revealed that this enzyme is localized at the
contact site between osteoclasts and the bone resorption
pit. Subsequently, it was shown that mutations in the gene
encoding cathepsin K are the cause of pycnodysostosis, a
rare hereditary disease which is characterized by hardened and fragile bones, short stature, skull deformities,
and osteoclasts that demineralize bone normally but
do not degrade its protein matrix. Evidently, cathepsin
K functions to degrade the protein matrix of bone and
hence is an attractive drug target for the treatment of
osteoporosis.
c. SARs and QSARs Are Useful Tools for
Drug Discovery
A lead compound is used as a point of departure to design more efficacious compounds. Experience has shown
that even minor modifications to a drug candidate can result in major changes in its pharmacological properties.
Thus, one might place methyl, chloro, hydroxyl, or benzyl
groups at various places on a lead compound in an effort
to improve its pharmacodynamics. For most drugs in use
today, 5 to 10 thousand related compounds were typically
synthesized in generating the medicinally useful drug.
These were not random procedures but were guided by experience as medicinal chemists tested various derivatives
of a lead compound: For those compounds that had
improved efficacy, derivatives were made and tested; etc.
This process has been systematized through the use of
structure–activity relationships (SARs): the determination, via synthesis and screening, of which groups on a lead
compound are important for its drug function and which
are not. For example, if a phenyl group on a lead compound interacts hydrophobically with a flat region of its
receptor, then hydrogenating the phenyl ring to form a
nonplanar cyclohexane ring will yield a compound with reduced affinity for the receptor.
A logical extension of the SAR concept is to quantify
it, that is, to determine a quantitative structure–activity relationship (QSAR). This idea is based on the premise that
there is a relatively simple mathematical relationship between the biological activity of a drug and its physicochemical properties. For instance, if the hydrophobicity of
a drug is important for its biological activity, then changing
the substituents on the drug so as to alter its hydrophobicity will affect its activity. A measure of the substance’s
hydrophobicity is its partition coefficient, P, between the
two immiscible solvents, octanol and water, at equilibrium:
P
concentration of drug in octanol
concentration of drug in water
[15.15]
Biological activity may be expressed as 1C, where C is the
drug concentration required to achieve a specified level of
biological function (e.g., IC50). Then a plot of log 1C versus log P (the use of logarithms keeps the plot on a manageable scale) for a series of derivatives of the lead compound having a relatively small range of log P values often
indicates a linear relationship (Fig. 15-29a), which can
therefore be expressed:
(a)
1
log __
C
1
log a b k1 log P k2
C
log P
(b)
Here k1 and k2 are constants, whose optimum values in this
QSAR can be determined by computerized curve-fitting
methods. For compounds with a larger range of log P values, it is likely that a plot of log 1C versus log P will have
a maximum value (Fig. 15-29b) and hence be better described by a quadratic equation:
1
log a b k1 1log P2 2 k2 log P k3
C
1
log __
C
log P
Hypothetical QSAR plots of log(1/C) versus
log P for a series of related compounds. (a) A plot that is best
described by a linear equation. (b) A plot that is best described
by a quadratic equation.
[15.16]
[15.17]
Of course, the biological activities of few substances
depend only on their hydrophobicities. A QSAR can
therefore simultaneously take into account several physicochemical properties of substituents such as their pK values, van der Waals radii, hydrogen bonding energy, and
conformation. The values of the constants for each of the
terms in a QSAR is indicative of the contribution of that
term to the drug’s activity. The use of QSARs to optimize
the biological activity of a lead compound has proven to
be a valuable tool in drug discovery.
FIGURE 15-29
d. Structure-Based Drug Design
Since the mid 1980s, dramatic advances in the speed and
precision with which a macromolecular structure can be
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Section 15–4. Drug Design
determined by X-ray crystallography and NMR (Section
8-3A) have enabled structure-based drug design, a process
that greatly reduces the number of compounds that need
be synthesized in a drug discovery program. As its name
implies, structure-based drug design (also called rational
drug design) uses the structure of a receptor in complex
with a drug candidate to guide the development of more
efficacious compounds. Such a structure will reveal, for example, the positions of the hydrogen bonding donors and
acceptors in a receptor binding site as well as cavities in
the binding site into which substituents might be placed on
a drug candidate to increase its binding affinity for the receptor. These direct visualization techniques are usually
supplemented with molecular modeling tools such as the
computation of the minimum energy conformation of a
proposed derivative, quantum mechanical calculations that
determine its charge distribution and hence how it would
interact electrostatically with the receptor, and docking
simulations in which an inhibitor candidate is computationally modeled into the binding site on the receptor to
assess potential interactions. Structure-based drug design
is an iterative process: The structure of the receptor in complex with a compound with improved properties is determined in an effort to further improve its properties.
e. Combinatorial Chemistry and High-Throughput
Screening
As structure-based methods were developed, it appeared that they would become the dominant mode of drug
discovery. However, the recent advent of combinatorial
chemistry techniques to rapidly and inexpensively synthesize large numbers of related compounds combined with
the development of robotic high-throughput screening
techniques has caused the drug discovery “pendulum” to
again swing toward the “make-many-compounds-and-seewhat-they-do” approach. A familiar example of combinatorial chemistry is the parallel synthesis of the large
number of different oligonucleotides on a DNA chip
(Section 7-6B). Similarly, if a lead compound can be synthesized in a stepwise manner from several smaller modules, then the substituents on each of these modules can
be varied in parallel to produce a library of related
compounds (e.g., Fig. 15-30).
A variety of synthetic techniques have been developed
that permit the combinatorial synthesis of thousands of related compounds in a single procedure. Thus, whereas investigations into the importance of a hydrophobic group
at a particular position in a lead compound might previously have prompted the individual syntheses of only the
ethyl, propyl, and benzyl derivatives of the compound, the
use of combinatorial synthesis would permit the generation of perhaps 100 different groups at that position. This
R2CHO
N
O
R1
O
O
O
B. Introduction to Pharmacology
The in vitro development of an effective drug candidate is
only the first step in the drug development process. Besides
causing the desired response in its isolated target receptor,
a useful drug must be delivered in sufficiently high concentration to this receptor where it resides in the human body
without causing unacceptable side effects.
a. Pharmacokinetics Is a Multifaceted Phenomenon
The most convenient form of drug administration is
orally (by mouth). In order to reach its target receptor, a
drug administered in this way must surmount a series of
formidable barriers: (1) It must be chemically stable in the
highly acidic (pH 1) environment of the stomach and must
not be degraded by the digestive enzymes in the gastrointestinal tract; (2) it must be absorbed from the gastrointestinal tract into the bloodstream, that is, it must pass
through several cell membranes; (3) it must not bind too
tightly to other substances in the body (e.g., lipophilic substances tend to be absorbed by certain plasma proteins and
by fat tissue; anions may be bound by plasma proteins,
mainly albumin; and cations may be bound by nucleic
acids); (4) it must survive derivatization by the battery of
enzymes, mainly in the liver, that function to detoxify
xenobiotics (foreign compounds), as discussed below (note
that the intestinal blood flow drains directly into the liver
via the portal vein, so that the liver processes all orally ingested substances before they reach the rest of the body);
(5) it must avoid rapid excretion by the kidneys; (6) it must
pass from the capillaries to its target tissue; (7) if it is targeted to the brain, it must cross the blood–brain barrier,
which blocks the passage of most polar substances; and
(8) if it is targeted to an intracellular receptor, it must pass
through the plasma membrane and, possibly, other intracellular membranes. The ways in which a drug interacts
with these various barriers is known as its pharmacokinetics. Thus, the bioavailability of a drug (the extent to
which it reaches its site of action, which is usually taken to
be the systemic circulation) depends on both the dose given
and its pharmacokinetics. Of course, barriers (1) and (2)
can be circumvented by injecting the drug [e.g., some forms
O
R3NH2
N
R1
would far more effectively map out the potential range of
the substituent and possibly identify an unexpectedly active analog. Interestingly, QSAR and computational techniques have been combined in the development of “virtual
combinatorial chemistry,” a procedure in which libraries of
compounds are computationally “synthesized” and “analyzed” to predict their efficacy, thereby again reducing the
number of compounds that must actually be synthesized
in order to generate an effective drug.
R2
R2
R1
531
H
N
N
H
R3
O
FIGURE 15-30 The combinatorial synthesis of
arylidene diamides. If ten different variants of
each R group are used in the synthesis, then 1000
different derivatives will be synthesized.
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Chapter 15. Enzymatic Catalysis
of penicillin (Fig. 11-25) must be injected because their
functionally essential -lactam rings are highly susceptible
to acid hydrolysis], but this mode of drug delivery is
undesirable for long-term use.
Since the pharmacokinetics of a drug candidate is as
important to its efficacy as is its pharmacodynamics, both
must be optimized in producing a medicinally useful drug.
The following empirically based rules, formulated by
Christopher Lipinski and known as Lipinski’s “rule of
five,” state that a compound is likely to exhibit poor
absorption or permeation if:
1. Its molecular mass is greater than 500 D.
2. It has more than 5 hydrogen bond donors (expressed
as the sum of its OH and NH groups).
3. It has more than 10 hydrogen bond acceptors (expressed as the sum of its N and O atoms).
4. Its value of log P is greater than 5.
Drug candidates that disobey Rule 1 are likely to have low
solubilities and to only pass through cell membranes with
difficulty; those that disobey Rules 2 and/or 3 are likely to
be too polar to pass through cell membranes; and those
that disobey Rule 4 are likely to be poorly soluble in aqueous solution and hence unable to gain access to membrane
surfaces. Thus, the most effective drugs are usually a compromise; they are neither too lipophilic nor too hydrophilic.
In addition, their pK values are usually in the range 6 to 8
so that they can readily assume both their ionized and unionized forms at physiological pH’s. This permits them to
cross cell membranes in their unionized form and to bind
to their receptor in their ionized form. However, since the
concentration of a drug at its receptor depends, as we saw,
on many different factors, the pharmacokinetics of a drug
candidate may be greatly affected by even small chemical
changes. QSARs and other computational tools have been
developed to predict these effects but they are, as yet,
rather crude.
b. Toxicity and Adverse Reactions Eliminate Most
Drug Candidates
The final criteria that a drug candidate must meet are
that its use be safe and efficacious in humans. Tests for
these properties are initially carried out in animals, but
since humans and animals often react quite differently to
a particular drug, the drug must ultimately be tested in humans through clinical trials. In the United States, clinical
trials are monitored by the Food and Drug Administration
(FDA) and have three increasingly detailed (and expensive) phases:
Phase I. This phase is primarily designed to test the
safety of a drug candidate but is also used to determine its
dosage range and the optimal dosage method (e.g., orally
vs injection) and frequency. It is usually carried out on a
small number (20–100) of normal, healthy volunteers, but
in the case of a drug candidate known to be highly toxic
(e.g., a cancer chemotherapeutic agent), it is carried out
on volunteer patients with the target disease.
Phase II. This phase mainly tests the efficacy of the
drug against the target disease in 100 to 500 volunteer patients but also refines the dosage range and checks for side
effects. The effects of the drug candidate are usually assessed via single blind tests, procedures in which the patient is unaware of whether he/she has received the drug
or a control substance. Usually the control substance is a
placebo (an inert substance with the same physical appearance, taste, etc., as the drug being tested) but, in the
case of a life-threatening disease, it is an ethical necessity
that the control substance be the best available treatment
against the disease.
Phase III. This phase monitors adverse reactions from
long-term use as well as confirming efficacy in 1000 to 5000
patients. It pits the drug candidate against control substances through the statistical analysis of carefully designed
double blind tests, procedures in which neither the patients
nor the clinical investigators evaluating the patients’ responses to the drug know whether a given patient has
received the drug or a control substance. This is done to
minimize bias in the subjective judgments the investigators
must make.
Currently, only about 5 drug candidates in 5000 that enter preclinical trials reach clinical trials. Of these, only one,
on average, is ultimately approved for clinical use, with
40% of drug candidates passing Phase I trials and 50%
of those passing Phase II trials (most drug candidates that
enter Phase III trials are successful). In recent years, the
preclinical portion of a drug discovery process has averaged 3 years to complete, whereas successful clinical trials have usually required an additional 7 to 10 years. These
successive stages of the drug discovery process are increasingly expensive, so that to successfully bring a drug
to market costs, on average, around $300 million.
The most time-consuming and expensive aspect of a
drug development program is identifying a drug candidate’s rare adverse reactions. Nevertheless, it is not an uncommon experience for a drug to be brought to market
only to be withdrawn some months or years later when it
is found to have caused unanticipated life-threatening side
effects in as few as 1 in 10,000 individuals (the search for
new applications of an approved drug and its postmarketing surveillance are known as its Phase IV clinical trials).
For example, in 1997, the FDA withdrew its approval of
the drug fenfluramine (fen),
CH3
CH2
CH
NH
CH2
CH3
CH3
CH2
CF3
Fenfluramine
C
NH2
CH3
Phentermine
which it had approved in 1973 for use as an appetite suppressant in short-term (a few weeks) weight loss programs.
Fenfluramine had become widely prescribed, often for ex-
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533
tended periods, together with another appetite suppressant, phentermine (phen; approved in 1959), a combination
known as fen-phen (although the FDA had not approved
of the use of the two drugs in combination, once it approves a drug for any purpose, a physician may prescribe
it for any other purpose). The withdrawal of fenfluramine
was prompted by over 100 reports of heart valve damage
in individuals (mostly woman) who had taken fen-phen for
an average of 12 months (phentermine was not withdrawn
because the evidence indicated that fenfluramine was the
responsible agent). This rare side effect had not been observed in the clinical trials of fenfluramine, in part because,
being an extremely unusual type of drug reaction, it had
not been screened for.
c. The Cytochromes P450 Metabolize Most Drugs
Why is it that a drug that is well tolerated by the majority of patients can pose such a danger to others?
Differences in reactions to drugs arise from genetic differences among individuals as well as differences in their
disease states, other drugs they are taking, age, sex, and environmental factors. The cytochromes P450, which function in large part to detoxify xenobiotics and participate in
the metabolic clearance of the majority of drugs in use,
provide instructive examples of these phenomena.
The cytochromes P450 constitute a superfamily of
heme-containing enzymes that occur in nearly all living organisms, from bacteria to mammals [their name arises from
the characteristic 450-nm peak in their absorption spectra
when reacted in their Fe(II) state with CO]. Humans express 100 isozymes (catalytically and structurally similar
but genetically distinct enzymes from the same organism;
also called isoforms) of cytochromes P450, mainly in the
liver but also in other tissues (its various isozymes are
named by the letters “CYP” followed by a number designating its family, an uppercase letter designating its subfamily, and often another number; e.g., CYP2D6). These
monooxygenases (Fig. 15-31), which in animals are embedded in the endoplasmic reticulum membrane, catalyze
reactions of the sort
RH O2 2H 2e ∆ ROH H 2O
The electrons (e) are supplied by NADPH, which passes
them to cytochrome P450’s heme prosthetic group via the
intermediacy of the enzyme cytochrome P450 reductase.
Here RH represents a wide variety of usually lipophilic
compounds for which the different cytochromes P450 are
specific. They include polycyclic aromatic hydrocarbons
[PAHs, frequently carcinogenic (cancer-causing) compounds that are present in tobacco smoke, broiled meats,
and other pyrolysis products], polycyclic biphenyls (PCBs,
which were widely used in electrical insulators and as plasticizers and are also carcinogenic), steroids (in whose syntheses cytochromes P450 participate; Sections 25-6A and
25-6C), and many different types of drugs. The xenobiotics
are thereby converted to a more water-soluble form, which
aids in their excretion by the kidneys. Moreover, the newly
generated hydroxyl groups are often enzymatically conju-
X-Ray structure of cytochrome P450CAM from
Pseudomonas putida showing its active site region. The heme
group, the Cys side chain that axially ligands its Fe atom, and
the enzyme’s lipophilic substrate thiocamphor are shown in
ball-and-stick form with N blue, O red, S yellow, Fe orange, and
the C atoms of the heme, its liganding Cys side chain, and the
thiocamphor green, cyan, and pale blue-green, respectively. The
bonds liganding the Fe are gray. [Based on an X-ray structure by
Thomas Poulos, University of California at Irvine. PDBid 8CPP.]
FIGURE 15-31
gated (covalently linked) to polar substances such as glucuronic acid (Section 11-1C), glycine, sulfate, and acetate,
which further enhances aqueous solubility. The many types
of cytochromes P450 in animals, which have different substrate specificities (although these specificities tend to be
broad and hence often overlap), are thought to have arisen
in response to the numerous toxins which plants produce,
presumably to discourage animals from eating them.
Drug–drug interactions are often mediated by cytochromes P450. For example, if drug A is metabolized by
or otherwise inhibits a cytochrome P450 isozyme that metabolizes drug B, then coadministering drugs A and B will
cause the bioavailability of drug B to increase above the
value it would have had if it alone had been administered.
This phenomenon is of particular concern if drug B has a
low therapeutic index. Conversely, if, as is often the case,
drug A induces the increased expression of the cytochrome
P450 isozyme that metabolizes it and drug B, then coadministering drugs A and B will reduce drug B’s bioavailability, a phenomenon that was first noted when certain
antibiotics caused oral contraceptives to lose their efficacy.
Moreover, if drug B is metabolized to a toxic product, its
increased rate of reaction may result in an adverse reaction. Environmental pollutants such as PAHs or PCBs are
also known to induce the expression of specific cytochrome
P450 isozymes and thereby alter the rates at which certain
drugs are metabolized. Finally, some of these same effects
may occur in patients with liver disease, as well as arising
from age-based, gender-based, and individual differences
in liver physiology.
Although the cytochromes P450 presumably evolved to
detoxify and/or help eliminate harmful substances, in sev-
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O
O
HO
C
H
N
C
CH3
C
CH3
N
cytochrome P450
O2
N
The metabolic
reactions of acetaminophen that
convert it to its conjugate with
glutathione.
FIGURE 15-32
O
CH3
spontaneous
H2O
H2O
OH
O
OH
Acetaminophen
(N-acetyl-p-aminophenol)
Acetimidoquinone
SH
COO–
+
H3N
CH
CH2 O
O
CH2
CH2
C
NH
CH
C
NH
CH2
COO–
CH2
COO–
Glutathione
(-L-Glutamyl-L-cysteinyl-glycine)
O
C
H
CH3
N
S
COO–
+
H3N
CH
CH2
O
CH2
C
OH
NH
CH2 O
CH
C
NH
Acetaminophen–glutathione conjugate
eral cases they have been shown to participate in converting relatively innocuous compounds to toxic agents. For
example, acetaminophen (Fig. 15-32), a widely used analgesic and antipyretic (fever reducer), is quite safe when
taken in therapeutic doses (1.2 g/day for an adult) but in
large doses (10 g) is highly toxic. This is because, in therapeutic amounts, 95% of the acetaminophen present is enzymatically glucuronidated or sulfated at its ¬ OH group
to the corresponding conjugates, which are readily excreted. The remaining 5% is converted, through the action
of a cytochrome P450 (CYP2E1), to acetimidoquinone
(Fig. 15-32), which is then conjugated with glutathione, a
tripeptide with an unusual -amide bond that participates
in a wide variety of metabolic processes (Section 26-4C).
However, when acetaminophen is taken in large amounts,
the glucuronidation and sulfation pathways become saturated and hence the cytochrome P450-mediated pathway
becomes increasingly important. If hepatic (liver) glutathione is depleted faster than it can be replaced, acetimidoquinone, a reactive compound, instead conjugates with
the sulfhydryl groups of cellular proteins, resulting in often fatal hepatotoxicity.
Many of the cytochromes P450 in humans are unusually polymorphic, that is, there are several common alleles
(variants) of the genes encoding these enzymes. Alleles
that cause diminished, enhanced, and qualitatively altered
rates of drug metabolism have been characterized for many
of the cytochromes P450. The distributions of these various alleles differ markedly among ethnic groups and hence
probably arose to permit each group to cope with the toxins in its particular diet.
Polymorphism in a given cytochrome P450 results in
differences between individuals in the rates at which they
metabolize certain drugs. For instance, in cases that a cytochrome P450 variant has absent or diminished activity,
otherwise standard doses of a drug that the enzyme normally metabolizes may cause the bioavailability of the drug
to reach toxic levels. Conversely, if a particular P450
enzyme has enhanced activity (usually because the gene
encoding it has been duplicated one or more times), higher
than normal doses of a drug that the enzyme metabolizes
would have to be administered to obtain the required therapeutic effect. However, if the drug is metabolized to a
toxic product, this may result in an adverse reaction.
Several known P450 variants have altered substrate specificities and hence produce unusual metabolites, which also
may cause harmful side effects.
Experience has amply demonstrated that there is no
such thing as a drug that is entirely free of adverse reactions.
However, as the enzymes and their variants that participate in drug metabolism are characterized and rapid and
inexpensive genotyping methods are developed, it may be-
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Section 15–4. Drug Design
come possible to tailor drug treatment to an individual’s
genetic makeup rather than to the population as a whole.
C. HIV Protease and Its Inhibitors
Acquired immunodeficiency syndrome (AIDS), the only
major epidemic attributable to a previously unknown
pathogen to appear in the 20th century (it was first described in 1981), is caused by human immunodeficiency
virus type 1 (HIV-1; the closely related HIV-2, which we
shall not explicitly discuss here, also causes AIDS and has
a similar response to drugs). HIV-1, which was discovered
in 1983, is a retrovirus, a family of viruses that were independently characterized in 1970 by David Baltimore and
Howard Temin. The retroviral genome is a single-stranded
RNA that reproduces inside its host cell by transcribing
the RNA to double-stranded DNA in a process mediated
by the virally encoded enzyme reverse transcriptase
(Section 30-4C). The DNA is then inserted into the host
cell’s chromosomal DNA by a viral enzyme named
integrase and is passively replicated along with the cell’s
DNA. However, under activating conditions (which for
HIV-1 often is an infection by another pathogen), the
retroviral DNA is transcribed, the proteins it encodes are
expressed and inserted in or anchored to the host cell
plasma membrane, and new virions (virus particles) are
produced by the budding out of a viral protein-laden segment of plasma membrane so as to enclose viral RNA (Fig.
15-33).
HIV-1 is targeted to and specifically replicates within
helper T cells, essential components of the immune system
(Section 35-2A). Unlike most types of retroviruses, HIV-1
eventually kills the cells producing it. Although the helper
T cells within which HIV-1 are actively replicating are often destroyed by the immune system, those within which
the HIV-1 is latent (its DNA is not being transcribed) are
not detected by the immune system and hence provide a
reservoir of HIV-1 (other types of cells also harbor HIV-1).
Consequently, over a several year period after the initial
535
infection (during most of which the host exhibits no
obvious symptoms), the host’s immune system is steadily
depleted until it has deteriorated to the point that the host
regularly falls victim to and is eventually killed by opportunistic pathogens that individuals with normally functioning immune systems can readily withstand. It is this
latter stage of an HIV infection that is called AIDS. In the
absence of effective therapy, AIDS is almost invariably fatal. Through the year 2002, an estimated 30 million people
had died of AIDS and an estimated 42 million others,
largely in sub-Saharan Africa, were HIV-positive, numbers
that are increasing at the rate of 5 million per year. As a
consequence of this global catastrophe, HIV has been characterized and effective countermeasures against it have
been devised faster than for any other pathogen in history.
a. Reverse Transcriptase Inhibitors Are Only
Partially Effective
The first drug to be approved by the FDA (in 1987)
to fight AIDS was 3-azido-3-deoxythymidine (AZT;
zidovudine),
HOCH2
H
_
N
T
O
H
H
H
+
N
N
H
3-Azido-3-deoxythymidine
(AZT; zidovudine)
which had first been synthesized in 1964 as a possible anticancer agent (it was ineffective). AZT is a nucleoside analog that, on enzymatic conversion to its triphosphate in the
cell (the plasma membrane is impermeable to nucleoside
triphosphates), inhibits HIV-1 reverse transcriptase, as do
the several other drugs (Section 30-4C) that the FDA had
approved to treat AIDS prior to 1996. Unfortunately, these
agents only slow the progression of an HIV infection but
do not stop it. This is in part because they are toxic, mainly
FIGURE 15-33 The assembly, budding,
and maturation of HIV-1. SU is the
surface glycoprotein gp120 and TM is
the transmembrane protein gp41.
[After Turner, B.G. and Summers, M.F.,
J. Mol. Biol. 285, 4 (1999).]
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(a)
(a)
I
MA
II III
CA
p1
IV
NC
p6
gag
I
MA
II III
CA
p1
V
TF
VI
VII
PR
RT
VIII
RN
IN
gag–pol
(b)
Cleavage
site
HIV-1 polyproteins. (a) The organization of
the HIV-1 gag and gag–pol polyproteins. The symbols used are
MA, matrix protein; CA, capsid protein; NC, nucleocapsid
protein; TF, transmembrane protein; PR, protease; RT, reverse
transcriptase; RN, ribonuclease; and IN, integrase. (b) The
sequences flanking the HIV-1 protease cleavage sites (red
bonds) indicated in Part a.
FIGURE 15-34
to the bone marrow cells that are blood cell precursors,
and hence cannot be taken in large doses. More important,
however, is that reverse transcriptase, unlike most other
DNA polymerases (Section 30-2A), cannot correct its mistakes and hence frequently generates mutations (about one
per 104 bp and, since the viral genome consists of 104 bp,
each viral genome bears, on average, one new mutation).
Consequently, under the selective pressure of an anti-HIV
drug such as AZT, the drug’s target receptor rapidly evolves
to a drug-resistant form.
b. HIV-1 Polyproteins Are Cleaved by HIV-1 Protease
HIV-1, as do other retroviruses, synthesizes its proteins
in the form of polyproteins, which each consist of several
tandemly linked proteins (Fig. 15-34). HIV-1 encodes two
polyproteins, gag (55 kD) and gag–pol (160 kD), which are
both anchored to the plasma membrane via N-terminal
myristoylation (Section 12-3B). These polyproteins are
then cleaved to their component proteins through the action of HIV-1 protease, but only after this enzyme has excised itself from gag–pol. This process occurs only after the
virion has budded off from the host cell and results in a
large structural reorganization of the virion (Fig. 15-33).
The virion is thereby converted from its noninfectious immature form to its pathogenic mature form. If HIV-1 protease is inactivated, either mutagenically or by an inhibitor,
the virion remains noninfectious. Hence HIV-1 protease is
an opportune drug target.
c. Aspartic Proteases and Their Catalytic Mechanism
HIV-1 protease is a member of the aspartic protease
family (also known as acid proteases), so called because
I
II
III
IV
V
VI
VII
VIII
Sequence
... Ser -Gln-Asn- Tyr — Pro - Ile - Val - Gln ...
... Ala -Arg- Val -Leu — Ala -Glu- Ala -Met ...
... Ala - Thr - Ile -Met — Met-Gln-Arg- Gly ...
... Pro - Gly -Asn- Phe — Leu-Gln- Ser -Arg ...
... Ser -Phe-Asn- Phe — Pro -Gln- Ile - Thr ...
... Thr -Leu-Asn- Phe — Pro - Ile - Ser - Pro ...
... Ala -Glu- Thr - Phe — Tyr - Val -Asp- Gly ...
... Arg- Lys - Ile -Leu — Phe -Leu-Asp- Gly ...
these enzymes all contain catalytically essential Asp
residues that occur in the signature sequence Asp–
Thr/Ser–Gly. Humans have several known aspartic proteases including pepsin, a digestive enzyme secreted by the
stomach (its specificity is indicated in Table 7-2) that functions at pH 1 and which was the first enzyme to be recognized (named in 1825 by T. Schwann); chymosin (formerly
rennin), a stomach enzyme, occurring mainly in infants,
that specifically cleaves a Phe–Met peptide bond in the
milk protein -casein, thereby causing milk to curdle, making it easier to digest (calf stomach chymosin has been used
for millennia to make cheese); cathepsins D and E, lysosomal proteases that function to degrade cellular proteins;
renin, which participates in the regulation of blood pressure and electrolyte balance (Fig. 15-35); and -secretase
(also known as memapsin 2), a transmembrane protein
common in brain that participates in cleaving A precursor protein to yield amyloid- protein (A), which is implicated in Alzheimer’s disease (Section 9-5B). In addition,
many fungi secrete aspartic proteases, presumably to aid
them in invading the tissues they colonize.
Eukaryotic aspartic proteases are 330-residue monomeric proteins. The X-ray structure of pepsin (Fig. 15-36a),
which closely resembles those of other eukaryotic aspartic
proteases, reveals that this croissant-shaped protein consists of two homologous domains that are related by
approximate 2-fold symmetry (although only about 25
residues in the core sheets of each domain are closely
related by this symmetry). Each domain contains a catalytically essential Asp in an analogous position. The Xray structures of enzyme–inhibitor complexes of various
aspartate proteases indicate that substrates bind in a
prominent cleft between the two domains that could ac-
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Section 15–4. Drug Design
1
Asp-Arg-Val-Tyr-Ile-His-Pro-Phe-His-Leu-Val-Ile-His13
Angiotensinogen
H2O
renin
1
Asp-Arg-Val-Tyr-Ile-His-Pro-Phe-His-Leu10
+
Val-Ile-His
Angiotensin I
H2O
angiotensin converting enzyme (ACE)
1
Asp-Arg-Val-Tyr-Ile-His-Pro-Phe8
+
His-Leu
Angiotensin II
Renin participation in blood pressure
regulation. Renin proteolytically cleaves the 13-residue
polypeptide angiotensinogen to the 10-residue polypeptide
angiotensin I. This latter peptide is then cleaved by angiotensin
converting enzyme (ACE) to the 8-residue polypeptide
angiotensin II, which, on binding to its receptor, induces
vasoconstriction and retention of Na and water by the
kidneys, resulting in increased blood pressure. Consequently
there have been considerable efforts to develop both renin and
ACE inhibitors for the control of hypertension (high blood
pressure), although as yet, only ACE inhibitors have been
approved as drugs.
FIGURE 15-35
537
What is the catalytic mechanism of eukaryotic aspartic
proteases? Proteolytic enzymes, in general, have three essential catalytic components:
1. A nucleophile to attack the carbonyl C atom of the
scissile peptide to form a tetrahedral intermediate (Ser 195
serves this function in trypsin; Fig. 15-23).
2. An electrophile to stabilize the negative charge that
develops on the carbonyl O atom of the tetrahedral intermediate (the H-bonding donors lining the oxyanion hole,
Gly 193 and Ser 195, do so in trypsin; Fig. 15-25).
3. A proton donor so as to make the amide N atom of
the scissile peptide a good leaving group (the imidazolium
group of His 57 in trypsin; Fig. 15-23).
commodate an 8-residue polypeptide segment in an extended sheetlike conformation. The active site Asp
residues are located at the base of this cleft (Fig. 15-36a).
Pepsin’s pH rate profile (Section 14-4) suggests that it
has two ionizable essential residues, one with pK L 1.1
and the other with pK L 4.7, which are almost certainly the
carboxyl groups of its essential Asp residues. At the pH of
the stomach, the Asp residue with pK 4.7 is protonated
and that with pK 1.1 is partially ionized. This suggests that
the ionized carboxyl group acts as a nucleophile to form
the putative tetrahedral intermediate. However, no covalent intermediate between an aspartic protease and its substrate has ever been detected.
The two active site Asp residues in eukaryotic aspartic
proteases are in close proximity and both appear to form
hydrogen bonds to a bridging water molecule that is present in several X-ray structures of eukaryotic aspartic proteases (Fig. 15-36b). This, together with a variety of enzymological and kinetic data, led Thomas Meeks to propose
(a)
(b)
FIGURE 15-36 X-Ray structure of pepsin. (a) Ribbon diagram
in which the N-terminal domain (residues 1–172) is gold, the
C-terminal domain (residues 173–326) is cyan, the side chains
of the active site Asp residues are shown in ball-and-stick form
with C green and O red, and the water molecule that is bound
by these Asp side chains is represented by a large red sphere.
The protein is viewed with the pseudo-2-fold axis relating core
portions of the two domains tipped from vertical toward the
viewer. (b) Enlarged view of the active site Asp residues and
their bound water molecule indicating the lengths (in Å) of
possible hydrogen bonds (thin gray bonds). The X-ray structures
of other aspartic proteases exhibit similar interatomic distances.
[Based on an X-ray structure by Anita Sielecki and Michael
James, University of Alberta, Edmonton, Canada. PDBid
4PEP.]
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Chapter 15. Enzymatic Catalysis
538
the following catalytic mechanism for aspartic proteases
(Fig. 15-37):
1. An active site Asp carboxylate group, acting as a
general base, activates the bound water molecule, the socalled lytic water, to nucleophilically attack the scissile peptide’s carbonyl C as an OH ion. Proton donation (general
acid catalysis) by the second, previously uncharged active
site Asp stabilizes the oxyanion that would otherwise form
in the resulting tetrahedral intermediate.
d. HIV-1 Protease Inhibitors Are Effective
Anti-AIDS Agents
HIV-1 protease differs from eukaryotic aspartic proteases in that it is a homodimer of 99-residue subunits.
Nevertheless, its X-ray structure (Fig. 15-38a), determined
independently in 1989 by Alexander Wlodawer, by Manual
Navia and Paula Fitzgerald, and by Tom Blundell, closely
resembles those of eukaryotic aspartic proteases. Thus,
HIV-1 protease has the enzymatically unusual property
that its single active site is formed by two identical sym-
2. The N atom of the scissile peptide is protonated by
the first Asp (general acid catalysis) resulting, through
charge rearrangement and proton transfer to the second
Asp (general base catalysis), in amide bond scission.
Aspartic proteases are inhibited by compounds with tetrahedral carbon atoms at a position mimicking a scissile peptide bond (see below). This strongly suggests that these
enzymes preferentially bind their transition states (transition state stabilization), thereby enhancing catalysis.
H
H
R
N
C
R
N
R
C
O
H
O
C
R
O
1
H
O
O
–O
H
– C
O
C
O
Asp
(a)
H
H
O
O
H
Asp
C
O
O
Asp
Michaelis complex
Asp
Tetrahedral intermediate
2
R
O
R
H
C
+
O
H
N
H
(b)
H
O
C
–O
O
Asp
C
Asp
Products
Catalytic mechanism of aspartic proteases.
(1) The nucleophilic attack of the enzyme-activated water
molecule (red) on the carbonyl carbon atom of the scissile
peptide bond (green) to form the tetrahedral intermediate. This
reaction step is promoted by general base catalysis by the Asp
on the right and general acid catalysis by the Asp on the left
(blue). (2) The decomposition of the tetrahedral intermediate
to form products via general acid catalysis by the Asp on the
right and general base catalysis by the Asp on the left.
FIGURE 15-37
X-Ray structure of HIV-1 protease.
(a) Uncomplexed and (b) in complex with its inhibitor
saquinavir (structural formula in Fig. 15-41). In each structure,
the homodimeric protein is viewed with its 2-fold axis of
symmetry vertical and is shown as a ribbon diagram with one
subunit gold and the other cyan. The side chains of the active
site Asp residues, Asp 25 and Asp 25, as well as the saquinavir
in Part b, are shown in ball-and-stick form with C green, N
blue, and O red. Note how the hairpin “flaps” at the top of
the uncomplexed enzyme have folded down over the inhibitor
in the saquinavir complex. Compare these structures with that
of the similarly viewed pepsin in Fig. 15-36a. [Part a based on
an X-ray structure by Tom Blundell, Birkbeck College, London,
U.K., and Part b based on an X-ray structure by Robert
Crowther, Hoffmann-LaRoche Ltd., Nutley, New Jersey.
PDBids (a) 3PHV and (b) 1HXB.]
FIGURE 15-38
O
See the Interactive Exercises
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Section 15–4. Drug Design
metrically arranged subunits. Quite possibly HIV-1 protease resembles the putative primordial aspartic protease
that, through gene duplication, evolved to form the eukaryotic enzymes (although HIV-1 protease is well suited
to the limited amount of genetic information that a virus
can carry).
Once the structure of HIV-1 protease became available,
intensive efforts were mounted in numerous laboratories
to find therapeutically effective inhibitors of this enzyme.
In this process, 200 X-ray structures and several NMR
structures have been reported of HIV-1 protease, its mutants, and the proteases of other retroviruses, both alone
and in their complexes with a great variety of inhibitors.
Hence, HIV-1 protease is perhaps the most exhaustively
structurally studied protein.
Comparison of the X-ray structure of HIV-1 protease
alone (Fig. 15-38a) with that of its complexes with polypeptidelike inhibitors (e.g., Fig. 15-38b) reveals that, on binding an inhibitor, the hairpin “flaps” covering the “top”
of the substrate-binding cleft move down by as much as
7 Å to enclose the inhibitor. Such an inhibitor binds to the
539
2-fold symmetric enzyme in a two-fold pseudosymmetric
extended conformation such that the inhibitor interacts
with the enzyme much like a strand in a sheet (Fig.
15-39). On the “floor” of the binding cleft, each signature
sequence (Asp 25–Thr 26–Gly 27) is located in a loop that
is stabilized by a network of hydrogen bonds similar to that
observed in eukaryotic aspartic proteases. The inhibitor interacts with the enzyme via a hydrogen bond to the active
site residue Asp 25. However, contrary to the case for eukaryotic aspartic proteases (Fig. 15-36b), no X-ray structure of an HIV-1 protease contains a water molecule within
hydrogen bonding distance of Asp 25 or Asp 25. On the
flap side of the binding cleft, the inhibitor interacts with
Gly 48 and Gly 48 and with a water molecule that is not
the attacking nucleophile but which mediates the contacts
between the flaps and the inhibitor backbone.
Although HIV-1 protease specifically cleaves the gag
and gag–pol polyproteins at a total of 8 sites (Fig. 15-34b),
these sites appear to have little in common except that their
immediately flanking residues are nonpolar and mostly
bulky. Indeed, binding studies indicate that HIV-1 pro-
Flap side
S2
S4
P2
P4
Ile 50
N
H
Gly 48
N
H
S3
P1
P3
Ile 50
N
H
Gly 48
C
C
O
H
O
H
H
N
C
N
H
O
H
N
C
C
N
H
N
H
H
N
Asp 29
O
Asp 29
N
H
O
O
O
C
Gly 27
O HO
H
N
OC
O
Asp 29
Gly 27
Asp 25
O
Asp 25
P3
P1
P2
P4
S3
S1
S2
S4
Arrangement of hydrogen bonds between HIV-1
protease and a modeled substrate. In the nomenclature used here,
polypeptide residues in one subunit are assigned primed numbers
to differentiate them from the residues of the other subunit;
substrate residues on the N-terminal side of the scissile peptide
bond are designated P1, P2, P3, … , counting toward the
N-terminus; substrate residues on its C-terminal side are
FIGURE 15-39
H
N
C
C
O
O
O
O
H
N
C
C
O
O
Gly 48
N
H
O
O
O
C
S1
designated P1, P2, P3, … , counting toward the C-terminus;
and the symbols S1, S2, S3, … , and S1, S2, S3, … , designate the
enzyme’s corresponding residue-binding subsites. The scissile
peptide bond is marked by arrows. [After Wlodawer, A. and
Vondrasek, J., Annu. Rev. Biophys. Biomol. Struct. 27, 257
(1998).]
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Chapter 15. Enzymatic Catalysis
O
H
N
R
C
N
CH
CH
N
N
C
N
H
H
Indinavir (CrixivanTM)
R
CH2
CH
C
N
H
O
R
O
Reduced Amide
H
N
HO
OH
R
CH
CH
CH
O
O
N
Peptide Bond
CH
N
•H2SO4
O
H
N
S
O
N
N
Nelfinavir (ViraceptTM)
Hydroxyethylene
CH
CH
N
S
C
CH
O
N
O
CH
Ph
OH
H
N
R
O
O
OH
H
CH3
N
N
O
H
Ph
R
•CH3SO3H
H
H
O
OH
NHtBu
OH
H
C
CH2
R
H
N
OH
Ph H
OH
N
S
Ritonavir (NorvirTM)
Dihydroxyethylene
OH
H
N
O
H
N
CH
CH
R
CH2
C
H
CH
R
N
N
NH2
Comparison of a normal peptide bond (top) to
a selection of groups (red) that are isosteres (stereochemical
analogs) of the tetrahedral intermediate in reactions catalyzed
by aspartic proteases.
FIGURE 15-40
H
OH
O H
•CH3SO3H
H
Saquinavir (InviraseTM)
H
tease’s specificity arises from the cumulative effects of the
interactions between the enzyme and the amino acids in
positions P4 through P¿4 . However, three of the peptides
cleaved by HIV-1 have either the sequence Phe-Pro or TyrPro, which are sequences that human aspartic proteases do
not cleave. Hence, HIV-1 protease inhibitors containing
groups that resemble either of these dipeptides would be
unlikely to inhibit essential human aspartic proteases.
An effective HIV-1 protease inhibitor should resemble
a substrate with its scissile peptide replaced by a group that
the enzyme cannot cleave. Such a group should, preferably, enhance the enzyme’s affinity for the inhibitor.
Mimics of the tetrahedral intermediate (Fig. 15-37), that
is, transition state analogs, are likely to do so. Consequently, a variety of such groups (Fig. 15-40) have been investigated in efforts to synthesize therapeutically effective
inhibitors of HIV-1 protease.
Although HIV-1 protease has high in vitro affinity for
its polypeptide-based inhibitors, these substances have
poor oral bioavailability (they are degraded by digestive
N
N
O
Hydroxyethylamine
NHtBu
O
O
O
O
N
O
NH2
OH
N
O
S
O
Ph
Amprenavir (AgeneraseTM)
Some HIV-1 protease inhibitors that are in
clinical use. Note that in addition to its generic (chemical)
name, each drug has a proprietary trade name, here in
parentheses, under which it is marketed.
FIGURE 15-41
proteases) and pharmacokinetics (they do not readily pass
through cell membranes.). Consequently, therapeutically
effective HIV-1 protease inhibitors must be peptidomimetics (peptide mimics), substances that sterically and
perhaps physically, but not chemically, resemble polypeptides. The use of peptidomimetics also permits conformational constraints to be imposed on a drug candidate that
would not be present in the corresponding polypeptide.
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Chapter Summary
As of early 2003, the FDA had approved six HIV-1 protease inhibitors (Fig. 15-41), the first of which, saquinavir,
was sanctioned in late 1995. These peptidomimetics have
IC50s against HIV in culture ranging from 2 to 60 nM but
have little or no activity against human aspartic proteases
(KI’s 10 M). They are the first drugs to clearly prolong
the lives of AIDS victims. Their development, in each case,
was a complex iterative process that required the design,
synthesis, and evaluation of numerous related compounds.
In several cases, these investigations capitalized on the
wealth of experience gained in developing peptidomimetic
inhibitors of the aspartic protease renin and in the resulting stockpiles of these compounds.
All the FDA-approved HIV-1 protease inhibitors initially cause a rapid and profound decline in a patient’s
plasma HIV load, which is often paralleled by immune
system recovery. However, as we saw with reverse transcriptase inhibitors, mutant forms of the protease that are
resistant to the inhibitor being used arise, usually within
4 to 12 weeks. Moreover, such a mutant protease is likely
to be resistant to other HIV-1 protease inhibitors, because
all of the HIV-1 protease inhibitors are targeted to the
same binding site. This has led to the use of combination
therapies in which an HIV-1 protease inhibitor is administered together with one, or more often, two reverse transcriptase inhibitors. This is because any virus that gains resistance to one drug in a regimen will be suppressed by the
other drug(s) in that regimen. In addition, the HIV-1 protease inhibitor ritonavir has been shown to be a potent inhibitor of the cytochrome P450 isoforms (CYP3A4,5,7)
that metabolize other protease inhibitors and hence is usually prescribed in low dosage as an adjunct to another pro-
541
tease inhibitor to improve the latter’s pharmacokinetics.
The plasma virus levels in many patients who were
placed on combination therapy rapidly became undetectable and have remained so for several years. This, however, does not constitute a cure: If drug therapy is interrupted, the virus will reappear in the plasma because
certain tissues in the body harbor latent viruses that are
unaffected by and/or inaccessible to drug therapy. Thus,
the presently available anti-HIV medications must be
taken for a lifetime.
Current anti-HIV therapies are by no means ideal. To
maximize their oral bioavailability, some of the different
drugs must be taken well before or after a meal but others must be taken with a meal. To minimize the probability of resistant forms of HIV arising, the bioavailability of
each drug must be maintained at a certain minimum level
and hence each drug must be taken on a rigid schedule.
Moreover, these drugs have significant side effects, mainly
fatigue, nausea, diarrhea, tingling and numbness with ritonavir, and kidney stones with indinavir. Consequently,
numerous AIDS patients fail to take their medications
properly, which greatly increases the likelihood that they
will develop resistance to these drugs and infect others with
drug-resistant viruses. Finally, HIV-1 protease inhibitors,
being complex molecules, are difficult to synthesize and
therefore are relatively expensive, so that in the developing countries in which AIDS is most prevalent, governments and most individuals cannot afford to purchase these
drugs, even if they were to be supplied at cost. It is
therefore important that anti-HIV therapies be developed
that are easy for patients to comply with, are inexpensive,
and ideally, will totally eliminate an HIV infection.
CHAPTER SUMMARY
1 Catalytic Mechanisms Most enzymatic mechanisms
of catalysis have ample precedent in organic catalytic reactions. Acid- and base-catalyzed reactions occur, respectively,
through the donation or abstraction of a proton to or from a
reactant so as to stabilize the reaction’s transition state complex. Enzymes often employ ionizable amino acid side chains
as general acid–base catalysts. Covalent catalysis involves nucleophilic attack of the catalyst on the substrate to transiently
form a covalent bond followed by the electrophilic stabilization of a developing negative charge in the reaction’s transition state. Various protein side chains as well as certain coenzymes can act as covalent catalysts. Metal ions, which are
common enzymatic components, catalyze reactions by stabilizing developing negative charges in a manner resembling
general acid catalysis. Metal ion–bound water molecules are
potent sources of OH ions at neutral pH’s. Metal ions also
facilitate enzymatic reactions through the charge shielding of
bound substrates. The arrangement of charged groups about
an enzymatic active site of low dielectric constant in a manner that stabilizes the transition state complex results in the
electrostatic catalysis of the enzymatic reaction. Enzymes catalyze reactions by bringing their substrates into close proximity in reactive orientations. The enzymatic binding of the
substrates in a bimolecular reaction arrests their relative motions resulting in a rate enhancement. The preferential enzymatic binding of the transition state of a catalyzed reaction
over the substrate is an important rate enhancement mechanism. Transition state analogs are potent competitive inhibitors because they bind to the enzyme more tightly than
does the corresponding substrate.
2 Lysozyme Lysozyme catalyzes the hydrolysis of
(1S4)-linked poly(NAG–NAM), the bacterial cell wall
polysaccharide, as well as that of poly(NAG). According to
the Phillips mechanism, lysozyme binds a hexasaccharide so
as to distort its D-ring toward the half-chair conformation of
the planar oxonium ion transition state. This is followed by
cleavage of the C1 ¬ O1 bond between the D- and E-rings as
promoted by proton donation from Glu 35. Finally, the resulting oxonium ion transition state is electrostatically stabilized by the nearby carboxyl group of Asp 52 so that the E-ring
can be replaced by OH to form the hydrolyzed product. The
roles of Glu 35 and Asp 52 in lysozyme catalysis have been
verified through mutagenesis studies. Similarly, structural and
binding studies indicate that strain is of major catalytic importance in the lysozyme mechanism. However, mass spectrometry and X-ray studies have demonstrated that the
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Chapter 15. Enzymatic Catalysis
lysozyme reaction proceeds via a covalent glycosyl–enzyme
intermediate involving Asp52 rather than by the noncovalently bound oxonium ion intermediate postulated by the
Phillips mechanism.
3 Serine Proteases Serine proteases constitute a widespread class of proteolytic enzymes that are characterized by
the possession of a reactive Ser residue. The pancreatically
synthesized digestive enzymes trypsin, chymotrypsin, and
elastase are sequentially and structurally related but have different side chain specificities for their substrates. All have the
same catalytic triad, Asp 102, His 57, and Ser 195, at their active sites. The differing side chain specificities of trypsin and
chymotrypsin depend in a complex way on the structures of
the loops that connect the walls of the specificity pocket, as
well as on the charge of the side chain at the base of the specificity pocket. Subtilisin, serine carboxypeptidase II, and ClpP
are unrelated serine proteases that have essentially the same
active site geometry as do the pancreatic enzymes. Catalysis
in serine proteases is initiated by the nucleophilic attack of
the active Ser on the carbonyl carbon atom of the scissile peptide to form the tetrahedral intermediate, a process that may
be facilitated by the formation of a low-barrier hydrogen bond
between Asp 102 and His 57. The tetrahedral intermediate,
which is stabilized by its preferential binding to the enzyme’s
active site, then decomposes to the acyl–enzyme intermediate
under the impetus of proton donation from the Asp 102polarized His 57. After the replacement of the leaving group
by solvent H2O, the catalytic process is reversed to yield the
second product and the regenerated enzyme. The Asp
102–His 57 couple therefore functions in the reaction as a proton shuttle. The active Ser is not unusually reactive but is ideally situated to nucleophilically attack the activated scissile
peptide. The X-ray structure of the trypsin–BPTI complex indicates the existence of the tetrahedral intermediate, whereas
X-ray structures of a complex of elastase with the heptapeptide BCM7 have visualized both the acyl–enzyme intermediate and the tetrahedral intermediate.
The pancreatic serine proteases are synthesized as zymogens to prevent pancreatic self-digestion. Trypsinogen is activated by a single proteolytic cleavage by enteropeptidase. The
resulting trypsin similarly activates trypsinogen as well as chymotrypsinogen, proelastase, and other pancreatic digestive
enzymes. Trypsinogen’s catalytic triad is structurally intact.
The zymogen’s low catalytic activity arises from a distortion
of its specificity pocket and oxyanion hole, so that it is unable
to productively bind substrate or preferentially bind the catalytic reaction’s transition state.
4 Drug Design Drugs act by binding to and thereby
modifying the functions of receptors. Many promising drug
candidates, which are known as lead compounds, have been
found by methods in which a large number of compounds are
tested for drug efficacy in an assay that is a suitable surrogate
of the disease/condition under consideration. Lead compounds are then chemically manipulated in the search for
compounds with improved drug efficacy. Structure–activity
relationships (SARs) and quantitative structure–activity
relationships (QSARs) are useful tools in this endeavor.
Structure-based drug design uses the X-ray and NMR structures of drug candidates in complex with their target proteins,
together with a variety of molecular modeling tools, to guide
the search for improved drug candidates. However, the advent
of combinatorial chemistry and high-throughput screening
procedures has extended the “make-many-compounds-andsee-what-they-do” approaches to drug discovery. In order to
reach their target receptors, drugs must have favorable pharmacokinetics, that is, they must readily traverse numerous
physical barriers in the body, avoid chemical transformation
by enzymes, and not be excreted too rapidly. Most useful drugs
are neither too lipophilic nor too hydrophilic so that they can
both gain access to the necessary membranes and pass through
them. Drug toxicity, dosage, efficacy, and the nature of rare
adverse reactions are determined through extensive and carefully designed clinical trials. Most drugs are metabolically
cleared through oxidative hydroxylation by one of the 100
cytochrome P450 isozymes. This permits the hydroxylated
drugs to be enzymatically conjugated to polar groups such as
glucuronic acid and glycine, which increases their rates of excretion by the kidneys. Drug–drug interactions are frequently
mediated by cytochromes P450. Polymorphisms among cytochromes P450 are often responsible for the variations
among individuals in their response to a given drug, including adverse reactions.
The formulation of HIV-1 protease inhibitors to control
HIV infections is one of the major triumphs of modern drug
discovery methods. HIV are retroviruses that attack specific
immune system cells and thereby degrade the immune system
over a period of several years to the point that it is no longer
able defend against opportunistic infections. HIV-1 protease
functions to cleave the polyproteins in immature HIV-1 virions that have budded out from a host cell, thus generating the
mature, infectious form. HIV-1 protease is an aspartic protease that, as eukaryotic aspartic proteases such as pepsin, uses
its two active site Asp residues to activate its bound lytic water molecule as the nucelophile that attacks and thereby
cleaves specific peptide bonds in the substrate polyprotein.
All of the FDA-approved peptidomimetic inhibitors of HIV1 protease cause a rapid and profound decrease in plasma HIV
levels, although they do not entirely eliminate the virus. They
are used in combination with reverse transcriptase inhibitors
to minimize the ability of the rapidly mutating HIV to evolve
drug-resistant forms.
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DRUG DISCOVERY
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HIV-1 PROTEASE AND OTHER ASPARTIC PROTEASES
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PROBLEMS
1. Explain why -pyridone is not nearly as effective a catalyst
for glucose mutarotation as is -pyridone. What about -pyridone?
2. RNA is rapidly hydrolyzed in alkaline solution to yield a
mixture of nucleotides whose phosphate groups are bonded to either the 2 or the 3 positions of the ribose residues. DNA, which
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Problems
lacks RNA’s 2 OH groups, is resistant to alkaline degradation.
Explain.
3. Carboxypeptidase A, a Zn2-containing enzyme, hydrolyzes the C-terminal peptide bonds of polypeptides (Section
7-1A). In the enzyme–substrate complex, the Zn2 ion is coordinated to three enzyme side chains, the carbonyl oxygen of the
scissile peptide bond, and a water molecule. A plausible model
for the enzyme’s reaction mechanism that is consistent with Xray and enzymological data is diagrammed in Fig. 15-42. What
are the roles of the Zn2 ion and Glu 270 in this mechanism?
545
4. In the following lactonization reaction,
C
C
R
R
O
O
OH
CH2
CH2
C
COO
R
R
R
R
the relative reaction rate when R CH 3 is 3.4 1011 times that
when R H. Explain.
*5. Derive the analog of Eq. [15.11] for an enzyme that catalyzes the reaction:
ABSP
Assume the enzyme must bind A before it can bind B:
E A B ∆ EA B ∆ EAB S EP
CO2–
CHR
O
Glu 270
C
NH R
O–
C
H
O
O
2+
H
Zn
attack of
water
CHR
C
NH R
O–
C
H
O–
O
Zn2+
H
8. Wolfenden has stated that it is meaningless to distinguish
between the “binding sites” and the “catalytic sites” of enzymes.
Explain.
10. In light of the information given in this chapter, why are
enzymes such large molecules? Why are active sites almost always located in clefts or depressions in enzymes rather than on
protrusions?
CO2–
O
7. Suggest a transition state analog for proline racemase that
differs from those discussed in the text. Justify your suggestion.
9. Explain why oxalate (OOCCOO) is an inhibitor of oxaloacetate decarboxylase.
Michaelis complex
Glu 270
6. Explain, in thermodynamic terms, why an “enzyme” that
stabilizes its Michaelis complex as much as its transition state does
not catalyze a reaction.
Tetrahedral intermediate
11. Predict the effects on lysozyme catalysis of changing Phe
34, Ser 36, and Trp 108 to Arg, assuming that this change does
not significantly alter the structure of the protein.
*12. The incubation of (NAG)4 with lysozyme results in the
slow formation of (NAG)6 and (NAG)2. Propose a mechanism
for this reaction. What aspect of the Phillips mechanism is established by this reaction?
13. How would the lysozyme binding affinity of the following
(1S4)-linked tetrasaccharide
scissle bond
scission
CO2–
CHR
O
C
H
R
C
O
O
H
Zn2+
Enzyme-product complex
FIGURE 15-42
NAG
O
H
NHCOCH3
compare with that of NAG–NAM–NAG–NAM? Explain.
H
+
O–
NAM
H
N
Glu 270
H
NAG
CH2OH
O
H
H
Mechanism of carboxypeptidase A.
14. A major difficulty in investigating the properties of the
pancreatic serine proteases is that these enzymes, being proteins
themselves, are self-digesting. This problem is less severe, however, for solutions of chymotrypsin than it is for solutions of
trypsin or elastase. Explain.
15. The comparison of the active site geometries of chymotrypsin and subtilisin under the assumption that their similarities have catalytic significance has led to greater mechanistic
understanding of both these enzymes. Discuss the validity of this
strategy.
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Chapter 15. Enzymatic Catalysis
16. Benzamidine (KI 1.8 105M) and leupeptin (KI 1.8 107M)
O
CH3C
O
Leu
Leu
NH
CH
CH
(CH2)3
NH
C
H2N
C
NH2+
H2N
Benzamidine
NH2+
Leupeptin
are both specific competitive inhibitors of trypsin. Explain their
mechanisms of inhibition. Design leupeptin analogs that inhibit
chymotrypsin and elastase.
17. Trigonal boronic acid derivatives have a high tendency to
form tetrahedral adducts. 2-Phenylethyl boronic acid
OH
CH2
CH2
B
OH
2–Phenylethyl boronic acid
is an inhibitor of subtilisin and chymotrypsin. Indicate the structure of these enzyme–inhibitor complexes.
18. Tofu (bean curd), a high-protein soybean product that is
widely consumed in China and Japan, is prepared in such a way
as to remove the trypsin inhibitor present in soybeans. Explain
the reason(s) for this treatment.
19. Explain why mutating all three residues of trypsin’s
catalytic triad has essentially no greater effect on the enzyme’s
catalytic rate enhancement than mutating only Ser 195.
20. Explain why chymotrypsin is not self-activating as is
trypsin.
21. Does Lipinski’s “rule of five” predict that a hexapeptide
would be a therapeutically effective drug? Explain.
22. The preferred antidote for acetaminophen overdose is
N-acetylcysteine. Explain why the administration of this substance, which must occur within 8 to 16 hours of the overdose, is
an effective treatment.
23. Why would the activation of HIV-1 protease before the
virus buds from its host cell be disadvantageous to the virus?
Explain.