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26
Chapter • Transferase Inhibitors
. Fig. 26.26 Pyrogallol 26.59, gallic acid 26.60,
or tropolone 26.61 bind to COMT with micromolar afnity. Tolcapone 26.62, entacapone 26.63,
nitecapone 26.64, or nebicapone 26.65 have strong
electron-withdrawing groups directly on or conjugated to the aromatic ring. These compounds are
nanomolar, competitive inhibitors of catecholamine.
The linking of two moieties, each analogous to catecholamine or adenosine with arigid ve-membered
tether (amide bond and double bond, red) affords
the nanomolar bisubstrate-analogue inhibitor 26.66
the sulfonium group. Presumably, this hydroxyl function
is deprotonated due to its proximity to the magnesium
ion, the sulfonium group, and the ammonium group of
Lys 144, increasing its nucleophilicity for the SN2-like
transfer of the methyl group from the positively charged
sulfur of SAM 26.57. The second, probably uncharged
phenolic OH group is involved in ahydrogen bond with
Glu 199. The crystal structure was determined with asubstrate-like inhibitor 26.58 in which the nucleophilicity of
the oxygen atom is very strongly suppressed by two electron-withdrawing nitro groups (. Fig.26.25). Methyl
transfer does not occur any longer.
Molecules with multiply hydroxylated aromatic rings
such as pyrogallol 26.59, gallic acid 26.60, or tropolone
26.61 show weak, micromolar afnity for the enzyme.
The introduction of strongly electron-withdrawing nitro
or carbonyl groups to the aromatic rings leads to asignificant increase in the afnity of these substrate-like inhibitors. The inhibitors tolcapone 26.62, entacapone 26.63,
nitecapone 26.64, or nebicapone 26.65 all have asubstitution pattern with anitro group in ortho-position to the
nucleophilic hydroxyl group and asecond electron-with-
drawing group in para-position (. Fig.26.26). Crystallographic studies of these derivatives have shown that
their nitro groups match that of 26.58, which is oriented
towards the SAM substrate (. Fig.26.25). The second
electron-withdrawing substituent is located where the
other nitro group of 26.58 is, and is oriented towards
the surrounding solvent. Tolcapone 26.62 was approved
in 1997 as aperipherally and centrally acting COMT inhibitor. Its therapeutic use has been severely limited due
to observed liver toxicity. Concomitant administration
of l-DOPA and entacapone 26.63, which acts predominantly in the periphery, has proved more benecial. It
has been on the market since 1998 and contributes to
abalanced level of l-DOPA.
All of these drugs compete with catecholamines for
amagnesium ion in the binding site. In recent years,
nanomolar bisubstrate inhibitors such as 26.66 have
been developed. They displace both the SAM cofactor
and catecholamine from the binding pocket. The original building blocks of the parent molecular model compounds can be seen in the bisubstrate inhibitors. The
crystal structure of one of these inhibitors is shown in
. • Blocking the Transfer of Farnesyl and Geranyl Anchors
. Fig. 26.27 Superposition of the crystal structures of COMT with
SAM 26.57 (magenta carbon atoms) and the catecholamine-like inhibitor 26.58 (green carbon atoms) with the bisubstrate inhibitor 26.66
(light blue carbon atoms). (7 https://sn.pub/JOQ0rm)
. Fig.26.27. Its binding geometry largely matches the
adenosine part of SAM on the nucleoside side and the
catecholamine side with the nitro aromatic ring. The
correct choice of the connecting bridge between the two
substrate-analogous parts of the molecule is crucial for
the binding afnity. Arigid ve-membered chain consisting of an amide group and an E-congured double bond
represents the optimum. Transitioning to amore exible
structure by hydrogenation of the double bond reduces
the binding afnity by afactor of 100. Lengthening the
chain with an additional member results in afurther 25fold reduction in binding afnity. Bisubstrate-analogue
inhibitors are expected to achieve higher selectivity for
their target enzymes. In the present case, the rigid and
geometrically strained linker between the two parts of
the molecule is responsible for the pre-organization necessary for the binding of the pharmacophoric groups.
This pre-organization of the ligand provides an advantage in binding to the receptor. Further development
must show whether such bisubstrate inhibitors have
achance of entering into drug development.
26.11 Blocking the Transfer of Farnesyl
and Geranyl Anchors
Kinases and phosphatases are not the only proteins
that undergo posttranslational during signal transduction. The spatial location of proteins is often essential
for their proper function in the cell. Some proteins need
26
Chapter • Transferase Inhibitors
. Fig. 26.28 Farnesyldiphosphate 26.67 binds to FTase and occu-
pies apart of the large catalytic site. The crystal structure of the enzyme with this substrate was determined (colored image: farnesyldiphosphate 26.67 dark-green). Geranylgeranyl groups 26.68 that have
an elongated isoprenyl chain (formulas: isoprenyl chain indicated
in red instead of a black chain in 26.67) are transferred by GGTase.
Trp 102β and Tyr 365β border the binding pocket in FTase and determine substrate selectivity. After binding the farnesyl substrate, the
peptide substrate 26.69 (gray) with its CAAX terminus diffuses into
the binding pocket. Catalyzed by an adjacent zinc ion coordinating
the cysteine residue of the substrate, the farnesyl residue is transferred
to the thiol group of the cysteine. The diphosphate group is nucleophilically displaced. Acrystal structure of the resulting product 26.70
(light-green) was also determined. It is shown in the colored image in
the center, superimposed onto the binary complex. The farnesyl residue must move “forward” in the pocket (green arrow). The resulting
product coordinates to the zinc ion. With its two aliphatic residues
A1 (here Ile) and A2 (here Val) of the CAAX motif, the tetrapeptide
moiety is recognized by the enzyme. The terminal methionine (X)
forms a hydrogen bond with the carboxylate group of Glu 167α.
(7 https://sn.pub/9qIkq2)
. • Blocking the Transfer of Farnesyl and Geranyl Anchors
. Fig. 26.29 Development of compounds for
the inhibition of FTase. Compound 26.71 represents acompetitive inhibitor for farnesyldiphosphate 26.67. Compounds 26.72–26.77
are inhibitors that bind competitively to the
tetrapeptide substrate, CAAX. Only some of
them (26.72–26.74, 26.76) use their functional
groups (e.g., imidazole rings) to block the zinc
ion in the catalytic site. Compounds 26.75 and
26.76 inhibit FTase without direct coordination to the Zn2+. Compound 26.77 blocks
FTase and GGTase equipotently
to be anchored to amembrane. In addition to examples
in which aportion of the polymer chain becomes immersed in the membrane, proteins are known that are
anchored to the membrane by an added farnesyl 26.67 or
geranylgeranyl 26.68 anchor. These hydrophobic anchors
consist of isoprenoid units (. Fig.26.28). They attach
to proteins via cysteine residues located near the C-ter-
minus. Three classes of prenylating enzymes are known:
the farnesyl transferases (FTases) and the geranylgera-
nyl transferasesI andII (GGTaseI andII). Substrates
of these catalysts include the GTPases of the Ras, Rab,
and Rho families, lamins, and the γ-subunit of G-protein
heterotrimers. For FTases and GGTases to attach aprenyl anchor, the substrate proteins must have aCAAX
sequence 26.69 at their C-terminus (. Fig.26.28). Here,
Cstands for the cysteine to which the prenyl group is
transferred, and A1 and A2 are usually aliphatic amino
acids. IfX is serine, methionine, glutamine, or alanine,
the protein will be prenylated by an FTase. Aleucine at
this position prefers aGGTase as acatalyst.
Meanwhile, more than 250 proteins that require the
posttranslational attachment of a prenyl tail for their
function have been discovered. Interest in these prenylating enzymes, especially FTases, began in the early 1990s.
It was observed that RAS proteins, which in amutated
form mediate apermanent growth signal in cancer, need
to be farnesylated. Only then are they active. If farnesylation is missing, RAS activity will be suppressed. After
Chapter • Transferase Inhibitors
26
the prenyl group has been transferred in the cytoplasm
to the cysteine three amino acids from the C-terminus,
the protein enters the endoplasmic reticulum. There,
the AAX tripeptide tail is proteolytically cleaved and
amethyl group is transferred to the C-terminus through
acarboxymethylation step. Finally, the prenylated protein is anchored to the cell inner membrane. FTases and
GGTases contain a zinc ion in their catalytic center,
which is coordinated by acysteine, aspartate, and histidine residue. First, the farnesyl or geranylgeranyl diphosphate anchor (26.67 or 26.68) diffuses into the large
funnel-shaped binding pocket of this enzyme. FTases
and GGTases form aheterodimer with abarrel-like architecture that is almost exclusively composed of helical
structural elements. FTase specically recognizes the
shorter substrate farnesyl diphosphate 26.67 because
the bottom of its binding pocket is conned by Trp 102β
and Tyr 365β. After successful binding of the prenyl substrate, the peptide chain with the tetrapeptidic C-terminal
CAAX of the protein to be prenylated diffuses into the
catalytic site. There, the prenyl substrate provides alarge
interaction surface for the incoming peptide substrate.
Next, the farnesyl chain must move towards the
peptide substrate for the actual transfer reaction. The
CAAX substrate occupies the fourth coordination site
on the zinc ion with the thiol group of its cysteine. It
also binds with its hydrophobic aliphatic side chain A2
into the preformed binding pocket of the enzyme. The
side chain A1 protrudes into the surrounding solvent.
In the structure shown in . Fig.26.28, a methionine
occupies the Xposition and the C-terminal carboxylate group forms ahydrogen bond with Gln 167α. The
prenyl group is then transferred to the peptide chain by
nucleophilic attack of the cysteine in the substrate on the
carbon atom next to the diphosphate group. Finally, the
prenylated product 26.70 diffuses out of the catalytic center. Interestingly, this is the rate-determining step. There
is evidence that anew substrate molecule is required to
displace the product from the enzyme. To do this, the
product molecule takes up anew position and binds to
aregion of the binding pocket through which it leaves
the reaction center.
According to the outlined reaction mechanism, different concepts for the development of inhibitors for this
enzyme have been pursued. The rst attempts were aimed
at competing with the binding of isoprenoid diphosphate. For example, the isoprenoid analog α-hydroxyfar-
nesylphophonic acid 26.71 occupies the binding pocket
similarly to farnesyl diphosphate and forms extensive
interactions with the enzyme as well as with the CAAX
peptide substrate. The second and most commonly used
strategy is to displace the peptide substrate from the
binding site. This goal can be achieved through the development of peptidomimetics. An example is l-739750
26.72, an ester prodrug that caused tumor regression in
rats without systemic toxicity (. Fig.26.29).
It has also been possible to completely abandon pep-
tide lead structures. Examples are R115777 (tipifarnib)
26.73 from Janssen Pharma or BMS-214662 26.74 from
Bristol-Myers Squibb. Both use their imidazole groups
to coordinate to the zinc ion. Compound 26.74 replaces
the isopropyl group of the peptide at position A1 with its
thiophene ring. The inhibitor uses its benzyl group for
the A2 position to mimic the side chain of the isoleucine.
With ABT-839 26.75, Abbott has found acompound
that does not coordinate to the zinc ion at all. It has
amethionine group at the end that is very similar to the
peptide tail in positionX of the natural substrate. Lonafarnib 26.76, atricyclic derivative, was developed at Schering-Plough; its urea group is directed to the binding site
through which the processed substrate leaves the binding
pocket. This inhibitor also blocks the enzyme without
coordinating to the zinc ion. Compounds 26.72–26.76 all
show aselectivity advantage for FTase. Merck has developed the nonpeptide structure 26.77, which is apotent
inhibitor of both FTase and GGTaseI. Of course, as
with COMT (Sect.26.10), astrategy can be pursued that
seeks to displace both substrates from the binding pocket
simultaneously. The bisubstrate-analog inhibitors suffer
from being very large in order to compete successfully
with the two large substrates.
Clinical studies on the nonpeptidic farnesyltransfer-
ase inhibitors 26.72–26.77 are not advanced enough to
be judged. Monotherapy with these inhibitors has been
rather disappointing, although very promising results
have been seen with tipifarnib 26.73 in breast cancer.
It remains to be seen whether FTase inhibitors will be
used as monotherapy in cancer treatment or whether
they will be used more effectively in combination with
other cytostatic and hormone drugs. However, in recent
years, anew eld of drug development has opened up
for FTase inhibitors. They appear to be potential lead
structures for the treatment of infectious diseases caused
by pathogenic microorganisms such as Plasmodium (malaria), Trypanosoma (African sleeping sickness and Cha-
gas disease), and Leishmania (leishmaniasis, kala-azar).
The causative agent of fungal diseases such as Candida
albicans can also be fought in this way. Obviously the
posttranslational prenylation of their proteins is an essential step in the lifecycles of these organisms. We can
hope that the sequence differences in the transferases are
adequately large compared to the human enzymes to develop selective compounds.
26.12 Synopsis
Proteins can be modied after translation in the ribo-
-
some by the attachment of groups such as phosphate,
methyl, or acetyl, and larger building blocks such as
prenyl or geranyl moieties or polypeptide chains such
as ubiquitin or SUMO.
. • Bibliography and Further Reading
Kinases transfer phosphate groups from ATP to the
-
hydroxyl groups of Ser, Thr, or Tyr residues or the im-
idazole group of His. This switches on the biochemi-
cal function of the phosphorylated protein substrates;
phosphatases can reverse this step by cleaving the
phosphate group off again from the phosphorylated
amino acid residue.
The more than 530 human kinases act as switches in
-
signaling cascades; thus, they seem very attractive as
putative drug targets. However, their substrate ATP is
present in high concentrations in cells, it is recognized
by multiple proteins often with other functions, and
Nature has established many processes involving ki-
nases redundantly as afailsafe. This makes selective
competitive inhibition of kinases at the ATP-binding
site adifcult task.
Kinases are rather exible proteins that adapt to their
-
substrates. The adenine moiety of ATP is recognized
by apeptide strand in the hinge region. Pockets are
found adjacent to the ATP binding site and they are
called front and back pocket. They are not involved in
ATP recognition but they can be exploited to endow
competitive inhibitors with the required selectivity.
Inhibitors are proled against the kinase family
-
(kinome) and exhibit either high selectivity against
individual members or show promiscuous binding to
larger groups on the phylogenetic kinase family tree.
Interestingly, introduction of inert metal centers that
expand the basic coordination architecture to attach
pharmacophoric groups can succeed in producing
highly selective compounds.
Imatinib and its follow-up compound nilotinib bind
-
to the inactive conformation of BCR-ABL kinase.
They represent acompletely new approach to can-
cer therapy: They cure chronic myeloid leukemia by
inhibiting the product of a misregulated gene. As-
ciminib is ahighly selective allosteric inhibitor that
mimics afatty acid residue in the myristoyl pocket
and stabilizes the kinase in aglobally inactive state.
The bump-and-hole method allows aspecic thera-
-
peutic validation of the biological relevance of atar-
get protein as well as the optimization of an inhibitor
class. Genetically, the target protein is modied in its
substrate specicity (e.g., a kinase at its gatekeeper
residue) and implemented into amodel organism.
Selective inhibition of this protein under in vivo con-
ditions is achieved via inhibitors that are adapted to
the modied binding site of the engineered protein.
Phosphatases remove phosphate groups from Ser,
-
Thr, Tyr, and His residues, thus, switching off the
biochemical function of asubstrate protein. Two cat-
alytically different enzyme classes are known, either
operating through nucleophilic attack of awater mol-
ecule, which is highly polarized by two adjacent metal
ions, or through the nucleophilic attack of acysteine
residue via apathway similar to that in cysteine prote-
ases. In both cases, the tetrahedral phosphorous atom
is nucleophilically attacked.
PTP-1B initially appeared to be an ideal target to treat
-
the metabolic syndrome because it involves dephosphorylation of the insulin receptor kinase. Potent
inhibitors of this target with challenging druggability could be developed; however, sufcient selectiv
ity with respect to another phosphatase, TCPTP,
failed. Knock-out mice were unable to survive when
the genes of both phosphatases are simultaneously
turned off. Asimilar life-threatening situation can be
anticipated with insufciently selective inhibitors.
The full-length phosphatase Shp2 is autoinhibited
-
by its N-SH2 domain in the absence of aphosphorylated substrate. This state can be stabilized by an
allosteric inhibitor that binds simultaneously to all
three domains and “glues” them together. This blocks
the formation of the enzymatically active form of the
phosphatase.
Catechol-O-methyl transferase is representative for
-
the family of methyl transferases using S-adenos-
yl-l-methionine as a cofactor for methyl transfer
via its sulfonium group. It transfers methyl groups
to catecholamines such as dopamine, adrenaline, or
noradrenaline.
Inhibition of the methyl transferase reaction is
-
achieved by introduction of strong electron-withdrawing groups, such as nitro groups, at the aromatic ring of the natural substrates, producing substrate-like inhibitors.
Farnesyl and geranylgeranyl transferases transfer pre-
-
nyl anchor groups onto protein substrates exhibiting
aCAAX sequence on their C-terminus. The phosphorylated prenyl anchor is attacked by the nucleophilic cysteine thiol group, which is further polarized
through the coordination to aneighboring zinc ion in
the catalytic center.
Inhibitors of farnesyl and geranylgeranyl transferases
-
bind competitively either to the CAAX peptide substrate or the prenyldiphosphate substrate binding site.
Some of them show strong peptidomimetic character
and involve coordination of the zinc ion. However,
completely nonpeptidic inhibitors have also been developed, some of which bind without zinc coordination.
Bibliography and Further Reading
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cial role of protein phosphorylation in cell signaling and its use as
targeted therapy, Int. J. Mol. Med., 40, 271–280 (2017)
B. M. Klebl and G. Müller, Second-generation Kinase Inhibitors, Ex-
pert Opin. Ther. Targets 9, 975–993 (2005)
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Chapter • Transferase Inhibitors
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H. Kubinyi and G. Müller, Eds., Chemogenomics in Drug Discovery.
A Medicinal Chemistry Perspective, Wiley-VCH, Weinheim (2004)
R. Lorenz, J. Wu, F. W. Herberg, S. S. Taylor, R. A. Engh, Drugging
the Undruggable: How Isoquinolines and PKA Initiated the Era
of Designed Protein Kinase Inhibitor Therapeutics, Biochemistry,
60, 3470–3484 (2021)
M. A. Fabian, W. H. Biggs etal., A Small Molecule-Kinase Interac-
tion Map for Clinical Kinase Inhibitors, Nat. Biotech 23, 329–336
(2005)
S. W. Cowan-Jacob, V. Guez, et al., Imatinib (STI571) Resistance in
Chronic Myelogenous Leukemia: Molecular Basis of the Under-
lying Mechanisms and Potential Strategies for Treatment, Mini-Re-
views in Medicinal Chemistry, 4, 285–299 (2004)
P. J. Alaimo, M. A. Shogren-Knaak and K. M. Shokat, Chemical Ge-
netic Approaches for the Elucidation of Signalling Pathways, Curr.
Opin. Chem. Biol., 5, 360–367 (2001)
M. J. Chen, J. E. Dixon, G. Manning, Genomics and evolution of pro-
tein phosphatases, Sci. Signal., 10, eaag1796 (2017)
J. P. Vainonen, M. Momeny, J. Westermarck, Druggable cancer phos-
phatases, Sci. Transl. Med., 13, eabe2967 (2021)
M. Köhn, Turn and Face the Strange: A New View on Phosphatases,
ACS Cent. Sci., 6, 467–477 (2020)
S. M. Stanford, N. Bottini, Targeting Tyrosine Phosphatases: Time to
End the Stigma, Trends Pharmacol. Sci., 38, 524–540 (2017)
L. Bialy and H. Waldmann, Inhibitors of Protein Tyrosine Phospha-
tases: Next-Generation Drugs? Angew. Chem. Int. Ed., 44, 3814–
3839 (2005)
M. J. Bonifacio, P. N. Palma, L. Almeida and P. Soares-da-Silva, Cate-
chol-O-methyltransferase and Its Inhibitors in Parkinson’s Disease,
CNS Drug Reviews, 13, 352–379 (2007)
C. L. Strickland and P. C. Weber, Farnesyl Protein Transferase: A Re-
view of Structural Studies, Curr. Op. Drug Discov. Develop., 2,
475–483 (1999)
K. T. Lane and L. S. Beese, Structural Biology of Protein Farnesyl-
transferase and Geranylgeranyltransferase TypeI, J. Lipid Res.,
47, 681–699 (2006)
R. Roskoski Jr., Properties of FDA-approved small molecule protein
kinase inhibitors: A 2024 update, Pharmacol. Res., 200, 107059
(2024)
Y. A. Puius etal., Identication of a Second Aryl Phosphate-binding
Site in Protein-tyrosine Phosphatase 1B: A Paradigm for Inhibitor
Design, Proc. Natl. Acad. Sci. USA, 94, 13420–13425 (1997)
B. G. Szczepankiewicz etal. Discovery of a Potent, Selective Protein
Tyrosine Phosphatase 1B Inhibitor Using a Linked-Fragment
Strategy, J. Am. Chem. Soc., 125, 4087–4096 (2003)
L. F. Iversen etal., Steric Hindrance as a Basis for Structure-Based
Design of Selective Inhibitors of Protein-Tyrosine Phosphatases,
Biochemistry, 40, 14812–14820 (2001)
Andrew P. Combs etal., Structure-Based Design and Discovery of
Protein Tyrosine Phosphatase Inhibitors Incorporating Novel Isothiazolidinone Heterocyclic Phosphotyrosine Mimetics, J. Med.
Chem., 48, 6544–6548 (2005)
D. P. Wilson etal., Structure-Based Optimization of Protein Tyrosine
Phosphatase 1B Inhibitors: From the Active Site to the Second
Phosphotyrosine Binding Site, J. Med. Chem., 50, 4681–4698
(2007)
C. Wiesmann etal., Allosteric inhibition of protein tyrosine phospha-
tase 1B, Nat. Struct. Biol. & Mol. Biol., 11, 730–737 (2004)
P. Hof etal., Crystal Structure of the Tyrosine Phosphatase SHP-2,
Cell, 92, 441–450 (1998)
J. G. Fortanet etal., Allosteric Inhibition of SHP2: Identication of a
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phatase as Potential Chemotherapeutic Agents for Glioblastoma:
A Minireview, ChemMedChem., ChemMedChem, 16, 777–787
(2021)
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chol-O-methyltransferase, Nature, 368, 354–358 (1994)
C. Lerner, B. Masjost etal., Bisubstrate Inhibitors for the Enzyme
Catechol-O-methyltransferase (COMT): Inuence of Inhibitor
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Oxidoreductase Inhibitors
Contents
27.1 Redox Reactions in Biological Systems Use Cofactors – 484
27.2 Chemotherapeutics for Cancer and Bacteria:
Dihydrofolate Reductase Inhibitors – 487
27.3 HMG-CoA Reductase Inhibitors: The Changing
Fate of Drug Development – 490
27.4 Hitting aMoving Target: Aldose Reductase Inhibitors – 496
27.5 11
27.6 The Cytochrome P450 Enzyme Family – 502
27.7 What Makes Slow and Fast Metabolizers Dierent? – 506
27.8 Blocking the Degradation of Neurotransmitters:
27.9 Cyclooxygenase: AKey Enzyme in Pain Sensation – 512
27.10 Synopsis – 518
β
-Hydroxysteroid Dehydrogenase – 500
Monoamine Oxidase Inhibitors – 508
Bibliography and Further Reading – 519
© The Author(s), under exclusive license to Springer-Verlag GmbH, DE, part of Springer Nature 2024
G. Klebe, Drug Design, https://doi.org/10.1007/978-3-662-68998-1_27
Chapter • Oxidoreductase Inhibitors
27
Chemical reactions that occur through the exchange of
electrons are called redox reactions. Typically, in bio-
chemical redox processes, the carbon atom changes its
oxidation state. In total, carbon can assume oxidation
states ranging from −4 to +4. In general, oxidations
convert derivatives with asignicant number of directly
bonded hydrogen atoms to derivatives with more contacts to nitrogen, oxygen, and sulfur. Since these bonds to
the above electronegative elements are usually associated
with the introduction of polar functional groups, redox
reactions exert adecisive inuence on the physicochem-
ical properties of the oxidized substances. For example,
water solubility is increased. This is important for the
elimination of xenobiotics. Cytochrome P450 enzymes,
alarge group of oxidizing enzymes, are involved in the
corresponding metabolic transformations. On the other
hand, reductions are also of vital importance for the
organism. In these reaction steps, reactive aldehydes or
ketones are converted into alcohols, which can then be
more easily conjugated and eliminated (Sect.8.1). Transition metals, which can assume avariety of oxidation
states, are predestined to serve as electron donors and
acceptors in redox reactions. In biological systems, one
transition metal, iron, is often used for this purpose. Once
incorporated into aprotoporphyrin scaffold, it exists in
apenta- or hexavalent coordination state and can assume
oxidation states between +2 and +4. It also participates
in complexes with sulfur. There it forms interesting multinuclear structures called iron–sulfur clusters. In addition to iron, copper also plays arole as amediator in
biochemical redox processes.
Nature uses cofactors for enzyme-catalyzed redox reactions. They are embedded in the specic environment
of aprotein and, shielded from the surrounding solvent,
carry out the electron or hydride ion transfer from the
group to be oxidized to the group to be reduced. Cofactors may be tightly bound to the protein. In these cases,
they are called prosthetic groups and do not leave the en-
zyme during the reaction. Other loosely bound cofactors
can be taken up by the protein like the substrate, chemically modied, and nally released. These cofactors must
be regenerated in another independent reaction for the
next redox reaction cycle.
The oxidoreductase class of enzymes will be discussed
in this chapter. They are involved in many electron transfer reactions and require electrons or hydrogen in the
form of hydride ions. These particles are transferred by
cofactors such as NAD(P)+ (nicotinamide adenine dinucleotide (phosphate)) or the avin nucleotides FMN
(avin mononucleotide) and FAD (avin adenine dinucleotide) and the aforementioned iron atom in the heme
group. Since these enzymes are also often involved in
processes that are causally related to the development of
pathological disease situations, many drug therapies are
aimed at inhibiting these enzyme systems.
27.1 Redox Reactions in Biological
Systems Use Cofactors
As mentioned above, enzymes use cofactors to transfer electrons or hydride ions in redox reactions. NAD+/
NADP+ 27.1 (nicotinamide adenine dinucleotide phos-
. Fig. 27.1 Many enzymatic redox reactions use NAD+/NADP+
27.1 (nicotinamide adenine dinucleotide, P stands for a phosphate
group attached to the ribose ring) and NADH/NADPH 27.2 as acofactor for the transfer of electrons and/or hydride ions. The cofactor
is made up of three components: the nicotinamide, which bears an attached ribose sugar (orange), the central diphosphate unit (violet) and
the adenosine moiety (green). There are two different derivatives for
27.1 and 27.2: one with aphosphate and one with ahydroxide group.
Upon oxidation, the positively charged nicotinamide moiety takes on
a hydride ion (red) at the 4-position; upon reduction the H
released from this position
−
ion is
. • Redox Reactions in Biological Systems Use Cofactors
. Fig. 27.2 Examples for an
oxidation reaction with malate
dehydrogenase (top) and for
areduction with homoserine
dehydrogenase (bottom). The
transformation of ahydroxyl
group into aketone function or
vice versa (red) is carried out in
both reactions
. Fig. 27.3 The stereochemically unambiguous transfer of ahydride
ion from the NADPH cofactor to the double bond of the substrate
being reduced is accomplished deep in the protein’s binding pocket.
Crystal structure determination of the enzyme dihydrofolate reductase
with bound dihydrofolic acid (DHF) and cofactor (NADPH) provided detailed information about the course of the reduction step. The
two reaction sites come spatially very close to one another in the structure. Ahydride ion is transferred from the 4-position of the reduced
phate) and NADH/NADPH 27.2 serve as acceptors and
donors of hydride ions (. Fig.27.1). This cofactor consists of three components: the nicotinamide with an attached ribose ring, the central diphosphate moiety, and
the adenosine moiety. The latter may carry aphosphate
moiety on the 2′-OH group and is then referred to as
NADP+/NADPH. The redox-active part is the nicotinamide moiety, apyridine derivative. During oxidation,
the positively charged NADP+ accepts ahydride ion at
the 4-position of the pyridine ring. In the reverse reaction, an H− is released from the same position. Atotal of
two electrons are transferred. NAD(P)+ is loosely bound
to the enzyme. It can be easily exchanged and regenerated
on another protein for asubsequent reaction cycle. Typical oxidation and reduction reactions that can occur in
adehydrogenase or reductase are shown in . Fig.27.2.
nicotinamide ring onto the neighboring double bond of the DHF substrate (violet line). (7 https://sn.pub/ZFSx22)
In the binding pocket of such an enzyme, there is direct contact between the group to be oxidized or reduced
and the nicotinamide ring. The binding site in these proteins is usually shielded from the aqueous solvent by
ahydrophobic group, aloop, or an amino acid lid. On
the one hand, this ensures that the stereochemistry of the
hydride transfer is unambiguous. On the other hand, access to protons must be excluded, otherwise the enzyme
would not be able to reduce the substrate, as elementary
hydrogen would be generated. The interaction geometry
for such reaction steps is shown for dihydrofolate reductase in . Fig.27.3. Although enzyme-catalyzed reactions
are generally reversible, and the direction of the reaction
will depend on the concentration of cofactors in the environment, NADP/H is, with few exceptions, involved in
reduction reactions. Oxidation reactions are almost exclu-