Protein Chemistry
DOI: 10.1002/anie.200501023
Protein Posttranslational Modifications: The Chemistry
of Proteome Diversifications
Christopher T. Walsh,* Sylvie Garneau-Tsodikova, and Gregory J. Gatto, Jr.
Angewandte
Chemie
Keywords:
amino acids · enzymes · protein
modifications · proteomics
C. T. Walsh et al.Reviews
7342 www.angewandte.org  2005 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim Angew. Chem. Int. Ed. 2005, 44, 7342 – 7372
1. Introduction
There are two major mechanisms for expanding the
coding capacity of the 6000 (yeast) to 30,000 (human) genes
in eukaryotic genomes to generate diversity in the corresponding
proteomes, the inventory of all proteins in a cell or
organism. Proteomes may be two to three orders of magnitude
more complex (> 1000000 molecular species of proteins)
than the encoding genomes would predict. The first
route of diversification of proteins is at the transcriptional
level, by mRNA splicing, including tissue-specific alternate
splicing.[1,2]
This is a central topic in RNA metabolism in
eukaryotic biology.
The second route to proteome expansion is the focus of
this Review: covalent posttranslational modification (PTM)
of proteins at one or more sites.[3]
As the name implies, these
are covalent modifications that occur after DNA has been
transcribed into RNA and translated into proteins. The
nascent or folded proteins, which are stable under physiological
conditions, are then subjected to a battery of specific
enzyme-catalyzed modifications on the side chains or backbones.
Proteome diversification by covalent modification
occurs in prokaryotes but is much more extensively encountered
in nucleated cells, both in terms of types of modifications
and frequency of occurrence. About 5% of the genomes
of higher eukaryotes can be dedicated to enzymes that carry
out posttranslational modifications of the proteomes.
Two broad categories of protein PTM occur (Scheme 1).
The first subsumes all enzyme-catalyzed covalent additions of
some chemical group, usually an electrophilic fragment of a
cosubstrate, to a side chain residue in a protein. The side chain
modified is usually electron rich, acting as a nucleophile in the
transfer. The second category of PTM is covalent cleavage of
peptide backbones in proteins either by action of proteases or,
less commonly, by autocatalytic cleavage. Limited proteolysis
to control location, activity, and lifetime of each protein in
intracellular and extracellular milieus is a central strategy for
the regulation of the composition and function of proteomes.
Protein covalent modifications can be sorted along several
axes. One is by the identity of the protein side chain modified;
15 of the 20 common proteinogenic amino acid side chains
[*] Prof. C. T. Walsh, Dr. S. Garneau-Tsodikova, Dr. G. J. Gatto, Jr.
Department of Biological Chemistry and Molecular Pharmacology
Harvard Medical School
Boston, MA 02115 (USA)
Fax: (+1)617-432-0348
E-mail: Christopher_walsh@hms.harvard.edu
The diversity of distinct covalent forms of proteins (the
proteome) greatly exceeds the number of proteins predicted by
DNA coding capacities owing to directed posttranslational
modifications. Enzymes dedicated to such protein modifications
include 500 human protein kinases, 150 protein phosphatases,
and 500 proteases. The major types of protein
covalent modifications, such as phosphorylation, acetylation,
glycosylation, methylation, and ubiquitylation, can be classified
according to the type of amino acid side chain modified,
the category of the modifying enzyme, and the extent of
reversibility. Chemical events such as protein splicing, green
fluorescent protein maturation, and proteasome autoactivations
also represent posttranslational modifications. An
understanding of the scope and pattern of the many posttranslational
modifications in eukaryotic cells provides insight
into the function and dynamics of proteome compositions.
From the Contents
1. Introduction 7343
2. Covalent Addition: The Main Acts 7345
3. Covalent Addition: The Supporting Cast 7354
4. Cataloguing the Posttranslational
Modification 7360
5. Multiple and Tandem Posttranslational
Modification of Proteins 7361
6. Reversible versus Irreversible
Posttranslational Modification 7362
7. Controlled Proteolysis 7364
8. Autocleavage and Peptide-Bond
Rearrangement 7365
9. Peptide Bond Rearrangement without
Autocleavage 7368
10. Conclusions 7369
Scheme 1. Two categories of posttranslational modifications of proteins:
1) covalent modification of a nucleophilic amino acid side chain
by an electrophilic fragment of a cosubstrate; 2) cleavage of a protein
backbone at a specific peptide bond.
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undergo such diversification (Table 1). Another classification
is that by the fragment of cosubstrate or coenzyme that is
enzymatically coupled to the protein and the concomitant
chemical nature of the protein modification. This catalogue
includes S-adenosylmethionine (SAM)-dependent methylation,
ATP-dependent phosphorylation, acetyl CoA dependent
acetylation, NAD-dependent ADP ribosylation,
CoASH-dependent phosphopantetheinylation, and phosphoadenosinephosphosulfate
(PAPS)-dependent sulfurylation.
A third axis of categorization of PTM is by the new
function enabled by the covalent addition. These include gain
in catalytic function of enzymes that have acquired tethered
biotinyl, lipoyl, and phosphopantetheinyl groups, changes of
subcellular address for proteins undergoing various lipid
modifications (prenylation, palmitoylation, glycosyl phosphatidylinositol
(GPI) anchor attachment), and targeting of the
modified protein for proteolytic destruction by ubiquitylation
to mark transport to lysosomes or proteasomes.
Christpher T. Walsh, born in 1944, majored
in biology at Harvard and completed his
PhD in biochemistry in the lab of Fritz
Lipmann at the Rockefeller Institute of
Medical Research. He was on the MIT
faculty (1972–1987), and since 1987 has
been at Harvard Medical School. He served
as Chair of the Dept. of Chemistry at MIT
(1982–1987) and of the Dept. of Biological
Chemistry and Molecular Pharmacology at
Harvard Medical School (1987–1995). His
research interests lie in enzyme and inhibitor
mechanisms and in the biosynthesis of nonribosomal
peptide antibiotics.
Sylvie Garneau, born in QuØbec, Canada,
received her BSc (1995) and MSc (1997) in
chemistry from the UniversitØ Laval, where
she worked under the supervision of Robert
ChÞnevert and PersØphone Canonne. She
completed her PhD in chemistry in January
2003 at the University of Alberta with
John C. Vederas on the studies of new
antimicrobial agents acting on bacterial cell
walls. She is currently a postdoctoral fellow
with Christopher T. Walsh at Harvard Medical
School, studying halogenation and pyrrole
formation various of natural products.
Gregory J. Gatto, Jr., born in 1972, majored
in chemistry at Princeton University, where
he worked under the direction of Martin
Semmelhack. In 2003, he received his MD
and PhD degrees from the Johns Hopkins
University School of Medicine. There, he
worked in the lab of Jeremy Berg on the
structural biology of peroxisomal targeting
signal recognition. He is currently an NIH
postdoctoral fellow in the lab of Christopher
T. Walsh at Harvard Medical School,
studying the biosynthesis of the macrolide
immunosuppressants.
Table 1: Posttranslational protein modifications at the side chains.[a]
Residue Reaction Example
Asp phosphorylation protein tyrosine phosphatases;
response regulators in twocomponent
systems
isomerization to isoAsp
Glu methylation chemotaxis receptor proteins
carboxylation Gla residues in blood coagulation
polyglycination tubulin
polyglutamylation tubulin
Ser phosphorylation protein serine kinases and
phosphatases
O-glycosylation notch O-glycosylation
phosphopantetheinylation fatty acid synthase
autocleavages pyruvamidyl enzyme formation
Thr phosphorylation protein threonine kinases/phos-
phatases
O-glycosylation
Tyr phosphorylation tyrosine kinases/phosphatases
sulfation CCR5 receptor maturation
ortho-nitration inflammatory responses
TOPA quinone amine oxidase maturation
His phosphorylation sensor protein kinases in twocomponent
regulatory systems
aminocarboxypropylation diphthamide formation
N-methylation methyl CoM reductase
Lys N-methylation histone methylation
N-acylation by acetyl, biotinyl,
lipoyl, ubiquityl
groups
histone acetylation; swinging-arm
prosthetic groups; ubiquitin;
SUMO (small ubiquitin-like
modifier) tagging of proteins
C-hydroxylation collagen maturation
Cys S-hydroxylation (S-OH) sulfenate intermediates
disulfide bond formation protein in oxidizing environments
phosphorylation PTPases
S-acylation Ras
S-prenylation Ras
protein splicing intein excisions
Met oxidation to sulfoxide Met sulfoxide reductase
Arg N-methylation histones
N-ADP-ribosylation GSa
Asn N-glycosylation N-glycoproteins
N-ADP-ribosylation eEF-2
protein splicing intein excision step
Gln transglutamination protein cross-linking
Trp C-mannosylation plasma-membrane proteins
Pro C-hydroxylation collagen; HIF-1a
Gly C-hydroxylation C-terminal amide formation
[a] No modifications of Leu, Ile, Val, Ala, Phe side chains are known. A
more extensive list can be found in reference [3].
C. T. Walsh et al.Reviews
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2. Covalent Addition: The Main Acts
The five most common types of covalent additions to
proteins are phosphorylation, acylation, alkylation, glycosylation,
and oxidation, which are catalyzed by dedicated PTM
enzymes (Scheme 2). The protein products obtained in this
manner make up subsets of the proteome of an organism: the
phosphoproteome, the acyl proteome, the alkyl proteome, the
glycoproteome, and the oxidized proteome. In turn, each of
these subproteomes can contain substantial diversity.
2.1. Protein Phosphorylation
The mammalian phosphoproteomes have phosphoSer
(pS), phosphoThr (pT) and phosphoTyr (pY) residues with
a split of about 90:10 (pS, pT/pY; Figure 1).[4]
Bacterial and
fungal phosphoproteomes will also have phosphoHis and
phosphoAsp residues derived from proteins in two-component
signal-transduction cascades.[5]
The pathogenic bacterium
Pseudomonas aeruginosa exhibits more than 60 such
pathways.[6]
The enzymes dedicated to protein phosphorylation are
among the largest class of PTM enzymes. This superfamily of
protein kinases have been termed the kinome, with over 500
members in the human kinome.[7]
If these acted on average on
only 20 different protein substrates, or different residues
within a smaller subset of proteins, 10000 distinct molecular
forms of phosphorylated proteins would be produced. This is
most probably a substantial underestimate of the true size of
phosphoproteomes of higher eukaryotes, in which phosphorylation
sites can be predicted but, as yet, cannot be completely
measured. For example, the enzymatic activity of Abl protein
kinase is modulated by phosphorylation at up to 11 distinct
residues (Tyr, Thr, Ser; Figure 2).
Introduction of the charged, dianionic tetrahedral phosphate
group induces altered conformations in local protein
microenvironments[8]
and is often paired with cationic arginine
side chains (Figure 3). These local reorganizations of
protein domains often create the architectural impetus for
signal initiation, for example, in the four parallel MAP kinase
pathways in eukaryotes and in the activation of many
membrane receptor tyrosine kinases during autophosphorylation.
The enzymatic conversion of a neutral OH side chain
to dianionic phosphate has proven to be such a useful
conformation switch in protein-domain restructuring that it
has evolved into a major recurring theme in eukaryotic
proteome diversity.
2.2. Protein Acylation
The most common acyl chains found in proteins that have
undergone posttranslational modifications are C2 (acetyl, e.g.
histone tail acetylations),[9]
C14 (myristoylation at glycine
N termini),[10]
and C16 (palmitoylated-S-Cys residues).[11]
The
8-kDa chain of the small protein ubiquitin[12]
(and congeners)
Scheme 2. Five major types of covalent additions to protein side
chains: phosphorylation, acylation, alkylation, glycosylation, oxidation.
Figure 1. Phosphorylated forms of amino acid side chains in proteins:
phosphoSer (pS); phosphoThr (pT); phosphoTyr (pY); phosphoHis
(pHis); phosphoAsp (pAsp).
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is enzymatically transferred as an acyl moiety by ubiquityl
ligases and chemically falls into this group of PTMs. The
biological consequences of the acylation of a given protein
with C2, C14, C16, or 8-kDa chains are vastly different.
2.2.1. e-N-Acetylation of Lysine
Acetylations of multiple lysine residues in histone Nterminal
tails or at the C-terminus of the p53 transcription
factor[13]
are viewed as an integral part of the epigenetic code
that controls selective gene transcription. The acetyl-group
donor is the primary metabolite acetyl CoA, and dedicated
histone acetyltransferase (HAT) isoenzymes select distinct
combinations of the e-NH2 groups of Lys side chains in the Nterminal
tails of histones: two Lys residues on histone H2A,
four on H2B, four on H3, and four on H4 (Figure 4). Given
two copies of each histone per octamer core in nucleosomes,
there are 14 ” 2 = 28 potential Lys side chains available for
acetylation. Yeast histones with up to 13 acetylations have
been reported,[14]
which reflects almost 50% posttranslational
modification. The combinatorial possibilities for differentially
acetylated histone tails in nucleosomes become astronomically
large.
The acetyl groups on Lys side chains convert potentially
cationic side chains into groups, thus altering the charge
distribution. The N-acetyl Lys group is also specifically
recognized by discrete protein domains, termed bromodomains
(Figure 5), which are embedded in transcription factors
and associated proteins. Thus, the acetylation state of histone
tails can regulate the recruitment of transcription factor
machinery that controls the initiation of transcription of the
genes in the region of chromatin covered by those nucleo-
somes.[14]
In Section 5, we describe how five Lys residues at
the C terminus of the transcription factor p53 can either be
acetylated or ubiquitylated. Acetylation blocks the Lys side
chains from ubiquitylation and prolongs the half-life of the
p53 protein molecule.
2.2.2. N-Myristoylation and S-Palmitoylation
Myristoylation of eukaryotic proteins at an N-terminal
glycine residue is catalyzed by the PTM enzyme protein Nmyristoyltransferase,
which utilizes the C14 myristoyl CoA as
the donor substrate.[15]
As protein synthesis is initiated with
N-terminal methionine residues, cotranslational hydrolysis of
the Met1–Gly2 bond by methionine aminopeptidase is a
prerequisite to myristoylation (Scheme 3). The newly generated
free amino group of the now N-terminal Gly is the
nucleophile in the acylation reaction. The introduced hydrophobic
C14 fatty acyl group can be a membrane-directing
Figure 2. Eleven phosphorylation sites in the tyrosine kinase Abl,
including Ser, Thr, and Tyr residues, color coded from red near the
N terminus to purple near the C terminus. This figure and all other
three-dimensional structural representations in this Review were
generated with MolScript.[122]
Figure 3. Conversion of a neutral OH side chain to a dianionic PO3
2À
side chain recruits cationic Arg side chains to make bifurcated chargepairing
interactions that can restructure the microenvironment of a
protein region, induce a conformational change, and thereby initiate or
propagate signaling information to partner proteins or small molecules.
Shown is the interaction in Cdk2 of pThr160 with the guanidinium
side chains of Arg50, Arg126, and Arg150.
Figure 4. There are 15 Lys residues in the N-terminal tails of H2A,
H2B, H3, and H4 that are sites for possible enzymatic acetylation. The
dark gray flags represent sites of acetylation on the indicated lysine (K)
residues.
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group to move proteins to membrane interfaces.[10]
Additionally
the myristoyl tail can switch from being buried in a cleft
within the protein in one state to being available for
membrane insertion in another conformational state.[16]
Examples of N-myristoylated proteins include HIV Gag
protein and protein kinase A.[17]
Palmitoylation of proteins requires the C16 fatty acyl CoA
as donor for the PTM acyltransferases, but the acyl group is
typically transferred to the sulfhydryl side chains of Cys
residues rather than to N-termini of proteins (Scheme 3).[11]
Among the most-well-studied examples are S-palmitoylation
of cysteines in the C-terminal regions of proteins such as the
Ras GTPase. The lipidation of these nucleophilic thiolate side
chains is consequential for partitioning Ras from the cytoplasm
to membrane interfaces to meet up with its signaling
partner proteins.[10]
As explained in Section 5, the S-palmitoylation
of Ras isoforms is part of a cascade of reversible
posttranslational modifications involved in maturation and
membrane anchoring of modified forms of Ras.[18]
Protein
substrates for S-palmitoylation can be transmembrane receptors
such as the CD8a chain and the CCR chemokine
receptor or cytoplasmic proteins such as the protein tyrosine
kinase Lck.[11]
2.2.3. Mono- and Polyubiquitylation of Proteins
An extension of the logic of posttranslational transfer of
low-molecular-weight (C2, C14, C16)
acyl chains to proteins is the acylation
on lysine e-amino groups by the
carboxy terminus of the 8-kDa protein
ubiquitin. In analogy to acyl
CoA donors of the electrophilic
acetyl, myristoyl, and palmitoyl acyl
groups, the C-terminal carboxy
group of the 76-residue ubiquitin
must be preactivated for acyl transfer.
The activation principle is the
same as that of the acyl thioester
system, but eukaryotic cells make
use of ubiquityl-S-protein as donors
instead of ubiquityl CoA.[19,20]
The
enzymatic activation machinery
involves an enzyme 1 to make ubiquityl-AMP
and a set of about a
dozen thiol-containing enzymes 2
(Scheme 4) that provide the activesite
nucleophile to capture the ubiquityl
group as a set of ubiquityl-Senzyme
2 family members. In general
a third set of proteins, known
collectively as enzyme 3 variants, are
then required to catalyze the efficient
transfer of the activated ubiquityl
protein tag to Lys side chains of
client proteins. Some of the
enzyme 3 subclasses are multicomponent
catalysts, with up to four
subunits, which provide selectivity
for a given protein target.[21]
There are several hundred
isoforms of such E3 ubiquityl ligases in higher eukaryotes,
which allow subtle discrimination among many target proteins
selected for ubiquitylation.[22]
Figure 5. The acetylation status of the histone N-terminal tail can
recruit partner proteins to control transcriptional activity on a nucleosome.
The interaction of an acetyl-e-NH-Lys side chain on a peptide
fragment of histone H3 with a bromodomain of coactivator protein is
redrawn from reference [32].
Scheme 3. a) Prior cleavage of the Met1–Gly2 peptide bond by methionine aminopeptidase liberates
the NH2 of the Gly residue for N-myristoylation by N-myristoyltransferase with myristoyl CoA as
donor. b) The thiolate side chain of a Cys residue as nucleophile towards palmitoyl CoA, catalyzed
by palmitoyl-S-protein transferases.
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Unlike the small-molecule acyl groups described in
Section 2.2.2., the ubiquityl acyl moiety provides an information-rich
architectural scaffold (Figure 6) that can be read by
particular partner proteins that control the downstream
biological response. Two types of protein
ubiquitylation can be distinguished according
to the number of ubiquitins added:
monoubiquitylation and polyubiquitylation.
Polyubiquitylation specifically refers to the
enzymatic construction of chains of ubiquitin
molecules. As shown in Figure 6a, ubiquitin
has Lys side chains on different
surfaces and tandem attachment could go
from any Lys unit on one ubiquitin monomer
to the C-terminal Gly76 on the preceding
monomer. It appears that polyubiquitin
chains are built up most often through Lys48
side chains, although chains tethered
through Lys63 are also well-known.[23]
The
X-ray structure of a Ub4 chain is shown in
Figure 6b. It is not clear how E3 ligases act
processively to build up Ubn chains (up to
Ub20) tethered to proteins in cells.
Monoubiquitylation and polyubiquitylation
consign proteins to relocation in cells,
most often with the net consequence of proteolytic degradation,
albeit by quite different mechanisms. Protein monoubiquitylation
at a Lys-e-NH2 group by an E3 ubiquitin ligase can
initiate relocation of transmembrane receptor proteins from
the plasma membrane to the trans
Golgi network sorting compart-
ments.[24–26]
The covalent Ub tag is
information-rich and recruits various
partner proteins that contain one or
more of several variants of Ub-binding
domains. The partner proteins
can act as chaperones for internalization
of the ubiquitylated receptor,
import into early endosomes, and
transit to lysosomes in which lysosomal
proteases can cause hydrolytic
degradation (Scheme 5).[23]
These
pathways are part of the homeostatic
regulation of receptor density and
lifetimes at plasma membranes.
In contrast to the sorting fate of
monoubiquitylated proteins, polyubiquitylation
sends modified proteins
to the chambered proteases that
constitute the proteasome. Tandem
attachment of four or more ubiquitin
molecules to such a lysine side chain
constitutes an architectural signal
that recruits protein chaperones
with ubiquitin-binding domains. The
chaperone complexes then escort the
marked protein to the protea-
somes,[27]
where the chaperones dissociate,
and the polyubiquitin chain is
removed hydrolytically, perhaps
during ATP-driven unfolding of the
targeted protein (Scheme 6). The
unfolded protein is then threaded
Scheme 4. Activation of the C-terminus of ubiquitin (at residue Gly76) by enzyme 1 to make Ub-AMP,
followed by transfer to a Cys thiolate in the active site of enzymes 2 to yield Ub-S-E2. Enzymes 3 can
act as chaperones and recruiters of specific proteins for ubiquitylation at Lys side chains.
Figure 6. a) 3D trace of the 76-residue ubiquitin: Lys29,48,63 side chains on different faces of ubiquitin offer
different surfaces for tandem conjugation of growing polyubiquityl chains; b) structure of a tetraubiquityl
unit, the minimum chain length to direct polyubiquitylated proteins to the proteasome.
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into the chamber of the proteasome where the active sites of
the protease subunits degrade it to small peptides. The
characteristic temporal control of proteins destroyed during
the cell cycle, such as the cyclin subunits of cyclin-dependent
protein kinases,[28,29]
is effected by the E1–E2–E3 ubiquitin
ligase machinery. The activity of various multisubunit E3
ligases can be controlled by posttranslational states of the
catalysts or of the target proteins, such as phosphorylation of
particular Ser and Thr residues.
2.3. Protein Alkylation
Alkyl substituents are attached regiospecifically to proteins
by posttranslational modification enzymes. The three
common alkyl groups transferred are the methyl (C1), for
example, in histone methylations of Lys and Arg side
chains,[30]
or the C15 and C20 isoprenyl (farnesyl and geranylgeranyl)
groups (Scheme 7).[31]
The small C1 and the large
hydrophobic C15 and C20
groups each serve to introduce
hydrophobicity but they
do so to very different
degrees.
2.3.1. N-Methylations
Whereas C-, O-, and Smethylations
of protein side
chains are known,[3]
the reactions
of most contemporary
interest are the N-methylations
of Lys and Arg side
chains, particularly on the
same histone tails that are
acetylated. Indeed, covalent
posttranslational N-methylation
of histone tails complements
acetylation as the
second main part of writing
and reading the histone code
(Figure 7). For example, 7 of
the first 36 residues, Arg2,17,26
and Lys4,9,27,36 of histone H3
are known to be methylated
Scheme 6. Recognition of Ubn-tagged protein for chaperoning to proteasomes where the Ubn tag is retrieved
by hydrolysis of the isopeptide link to the target protein; the target protein is unfolded and threaded into
the chamber of the proteasome.
Scheme 5. Recognition of the Ub architecture in a monoubiquitintagged
protein by partner proteins/chaperones that have one or
more ubiquitin-recognition domains for transit to the secretory
system.
Scheme 7. Alkyl groups transferred to protein side chains: the methyl
group from S-adenosylmethionine (SAM) is transferred most often to Lys
and Arg side chains (although O-, S-, and C-methylations of protein side
chains are known); the two isoprenyl units transferred by protein
prenyltransferases to Cys side chains are the C15 (farnesyl) and the C20
(geranylgeranyl) groups from the corresponding prenyl diphosphate
substrates.
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by a family of histone methyltransferases,[32]
several of which
are residue-specific. An additional layer of information
content and combinatoric complexity is enabled by the fact
that Lys-e-NH2 groups can be progressivly mono-, di-, or
trimethylated, again with distinct distributions shown by
different methyltransferases. Analogously both monomethyland
dimethylArg residues are observed (Scheme 8). The size
and hydrophobicity differences between monomethyl- and
trimethyl substituents on Lys side chains enable selective
recruitment of proteins involved in transcriptional control.
For example, trimethyl-Lys9 in H3 recruits partner protein
HP1 by binding to its chromodomain (Figure 8) as part of
transcription factor and coactivator protein complex assem-
blies.[14,32]
N-Acetylations and N-methylations of side chains in
histone tails can have opposite effects on gene transcriptional
silencing and activation. Histone H3 is just one of the four
histones, each present as a dimer, in the nucleosome core. The
balance of the four acetylations and seven methylations on
the tail of histone H3 give, on this subunit alone, 11 ” 2 = 22
sites for titration of transcriptional coactivator and corepressor
complexes.
2.3.2. Protein S-Prenylation
The C15 farnesyl and C20 geranylgeranyl lipid groups are
built up from C5 isoprenyl diphosphate primary metabolites
by iterative alkyl extension by action of CÀC bond-forming
enzymes.[33]
The farnesyl and geranylgeranyl-PP molecules
can be used for further isoprene elongation (e.g. C15
dimerization to squalene in the cholesterol biosynthetic
pathway) or they can be utilized as
electrophilic alkyl donors in posttranslational
protein prenylation
(Scheme 9).[34]
There are protein farnesyltransferases
and protein geranylgeranyltransferases,
which are ab heterodimers
that share a common a subunit.
The Ras GTPase superfamily have
members that can be prenylated on
Cys thiolate side chains, some with the
C15 and some with the C20 prenyl chain.
Typically, Ras family proteins that have
a CaaX motif at the C terminus are
farnesylated, when X is a small amino
acid such as Ala or Ser. When X is Leu
as in both Rac and RhoA GTPases,
then the cysteine residue (C) is geranylgeranylated
(Scheme 10a). The Rab
subfamily of GTPases, over 60 in
number, have two cysteine residues at
or near the C terminus, for example, in
a CCXX arrangement. Both cysteine
groups undergo posttranslational geraFigure
7. N-methylations can occur at Arg2,17,26 and Lys4,9,27,36 of histone H3. Lys18
and Lys27 can be acetylated and Ser10 and Ser28 phosphorylated.
Scheme 8. Progressive mono-, di-, and trimethylation of Lys side chains and mono- and dimethylation of
Arg side chains in histone N-terminal tail regions. SAH=S-adenosylhomocysteine.
Figure 8. N,N,N-Trimethyl-Lys9 of histone H3 as a ligand to
recruit protein HP1 through its chromo domain.
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nylgeranylation (Scheme 10b), thus introducing two C20 lipid
anchors.[35]
Rab proteins cycle between membrane vesicles in
the secretory system; escort proteins are used to control
location and cycling.[36,37]
Some proteins undergo both N- and S-acylation and some
combination of S-prenylation. All of these lipid anchors drive
the modified proteins to partition
more to membranes,
thus controlling subcellular
localizations.
2.4. Protein Glycosylation
Covalent glycosylation of
proteins is relatively rare in
prokaryotes and quite
common in eukaryotes. Cglycosylation,
O-glycosylation,
and N-glycosylation of
proteins are known, but Cglycosylation,
specifically
mannosylation of C2 of the
indole ring of tryptophan res-
idues,[38]
is quite rare.
2.4.1. N-Glycosylation
N-glycoproteins are both
more common and typically
more complex in structure
and architecture than O-glycoproteins
in eukaryotes.[39]
The branching glycan unit in Nglycoproteins
is preassembled on a lipid diphosphate scaffold
by a series of membrane-associated glycosyltransferases in
the endoplasmic reticulum.[40]
The assembled N-glycan unit
that serves as a substrate for the multisubunit oligosaccharyltransferase
is a tetradecasaccharyl-PP-dolichol substrate
Scheme 9. Mechanism for Cys S-isoprenylation by protein prenyltransferases.
Scheme 10. Prenylation reactions at the C termini of the Ras protein superfamily: a) farnesylation of the C terminus of Ras at CaaX (X=Ala, Ser).
Geranylgeranylation of Rac at CaaX (X=Leu); b) double geranylgeranylation of the CC carboxy terminus of Rab proteins.
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(Scheme 11).[41]
The side-chain atoms that are modified in Nglycoprotein
biogenesis are the carboxamide nitrogen atoms
of asparagine groups, almost always in the sequence Ser/ThrX-Asn.
The low nucleophilicity of the Asn CONH2 is thought
to be enhanced by hydrogen bonding to the Ser/Thr-OH side
chain but the initial glycan transfer step is still poorly
understood.
The initial tetradecasaccharyl chain Glc3Man9(GlcNAc)2
then undergoes a remarkable enzymatic hydrolytic trimming
and refashioning of the identity and linkages of the N-glycan
chains (Scheme 12). The first hydrolytic tailoring enzymes are
Scheme 11. The branched tetradecasaccharyl-PP substrate is the donor substrate in the Asn N-glycosylation reaction catalyzed by the
oligosaccharyltransferase that initiates N-glycoprotein modifications.
Scheme 12. Progressive trimming of the initial N-linked Glc3Man9(GlcNAc)2 glycan chain to Man9(GlcNAc)2 in the ER and then to the core
pentasaccharide Man3(GlcNAc)2 in the Golgi complex before being built back up to mature N-glycan chains found on N-glycoproteins that have
transited the secretory system.
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two glucosidases in the ER lumen which generate the
dodecasaccharyl GlcMan9(GlcNAc)2 N-linked proteins.
These are ligands for calnexin and calreticulin, protein
chaperones that recognize the dodecasaccharyl chain and
help refold the nascent glycoproteins as they are extruded
through the ER membrane into the lumen. As the remaining
Glc is hydrolyzed by the ER glucosidase, the chaperones lose
affinity for the undecasaccharyl-N-protein products. A UDPglucose
glycoprotein glucosyltransferase[42]
puts the Glc
residue back on the undecasaccharide unit to give the
calreticulin and calnexin chaperones another round of
assisted refolding. Thus, this is a refolding/protein-qualitycontrol
way station during glycoprotein secretion. If the
glycoprotein cannot be refolded after several cycles of
deglucosylation/reglucosylation, it is targeted for export
back to the cytoplasm. There it undergoes polyubiquitylation,
and proteasome-mediated degradation of the unfolded protein
by a glycoprotein-targeting E3 ligase as part of the ERaccelerated
degradation (ERAD) quality-control pathway
(Scheme 13).[42]
For Man9(GlcNAc)2–N-glycoproteins that have refolded
and passed into the Golgi compartment, six of the mannose
residues are then trimmed hydrolytically by mannosidases to
yield the Man3(GlcNAc)2 pentasaccharyl core found in all
mature N-glycoproteins. At this point the branched oligosaccharide
core is rebuilt back up to biantennary and triantennary
oligosaccharides characteristic of mature N-glycoproteins
present on the cell surface.[43]
The multiplicity of
glycosyltransferases in the Golgi can create enormous diversity
in the mature N-glycan chains.
Multiple Asn side chains can be glycosylated in a given
protein, and the identity of the glycan chains at each Asn
residue can vary depending on stochastic encounters with at
least 10 trimming and rebuilding enzymes during passage
through the ER and Golgi compartments.[41,43]
It has been
estimated that about a third of all proteins that enter
secretory pathways in eukaryotic cells may be N-glycosylated,
and so tens of thousands of glycoprotein variants may coexist
in eukaryotic cells. For example, 52 glyco forms of the prion
protein have been reported.[44,45]
2.4.2. O-Glycosylation
O-glycosyl chains in eukaryotic proteins are generally
shorter and less complex than those in N-glycoproteins. Many
proteins contain the monosaccharide GlcNAc[46]
that is put on
by a specific O-GlcNAc transferase and removed by a
corresponding hydrolase. Other proteins such as the signaling
protein Notch contain tri- and tetrasaccharides in the EGF
repeat domains (Figure 9).[47,48]
O-glycosylation is a crucial
part of the maturation of Notch during its transit through the
secretory pathway to the cell surface. The short O-linked
sugar chains are important in a variety of functional contexts,
from modulating transcription factor activity,[49]
to acting as
essential recognition elements in signaling by Notch at cell
surfaces.[50,51]
2.5. SÀS Bond Formation
Two main types of linkages serve to cross-link proteins, or
portions of proteins, covalently. By far the more common are
disulfide links from oxidation of cysteinyl residue thiolate side
chains.[52,53]
The cytoplasmic and nuclear compartments in
eukaryotic cells are reducing microenvironments, as reflected
in the 100:1 ratio of the redox-active tripeptide glutathione in
reduced (GSH) to oxidized (GSSG) state.[54]
The high
reducing ratio is maintained by the high levels of NAD(P)H
and enzymes, such as glutathione reductase and thioredoxin
reductase[3]
that use the reduction potential of NAD(P)H to
re-reduce disulfides in proteins that have become oxidized
(Scheme 14a). As proteins pass through the secretory pathway
in eukaryotic cells, the levels of total glutathione and
reduced nicotinamide coenzyme fall, compartments become
Scheme 13. Cycling of the oligosaccharyl chain between GlcMan9(GlcNAc)2-Asn
and Man9(GlcNAc)2-Asn in the ER from opposing
action of glucosidase and UDP-glucose glycoprotein glucosyltransferase.
The Glc-containing oligosaccharyl chain is recognized by the
chaperone proteins calreticulin and calnexin to help refold the nascent
N-glycoproteins that have been glycosylated and extruded cotranslationally
into the ER lumen. The glucosylation cycle is part of the
protein quality-control system for proteins secreted into the endoplasmic
reticulum.
Figure 9. The O-linked tetrasaccharide sialyl-a-2,3-Gal-x-1,4-GlcNAc-b-
1,3-fucosyl-Ser attached to the protein Notch during its transit through
the secretory pathway on the way to the cell surface.
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more oxidizing, and disulfide links predominate. The disulfide
bonds may stabilize protein architectures as proteins reach
cell outer surfaces or are excreted into extracellular spaces.
The formation of disulfide bonds in zymogen forms of
pancreatic protease as they are packaged into the oxidizing
microenvironment of zymogen granules is a prototypical case.
The mechanism for oxidation of protein dithiols to
disulfides typically involves oxidation of the electron-rich
thiolate side chains of Cys residues. One-electron oxidation
would yield thiyl radicals that could dimerize to the disulfides.
Alternatively, thiolate side chains can be oxygenated by a
variety of oxygen-derived oxidants (peroxide, hydroxyl radical)
to generate sulfenic acid (-SOH) side chains. Capture of
the sulfenate by a neighboring Cys-SÀ
generates disulfides.
Regeneration of the dithiol forms is mediated by thiol–
disulfide interchange using reduced glutathione or the lowmolecular-weight
dithiol protein thioredoxin (TSH). The
oxidized GSSG or TSST are recycled at the expense of
NADPH oxidation by thioredoxin reductase and glutathione
reductases.
The electron-rich thiolates and the thiyl radicals can be
captured by other oxidants and radicals, including CNO. Such
S-nitrosylation of thiolate side chains of cysteine residues is
documented in many proteins by radical species derived from
nitric oxide (Scheme 14b). These Cys-SNO moieties have
been proposed to be widespread in oxidative signaling
events.[55]
The second PTM cross-link strategy is nonoxidative and
involves transglutaminase catalysis in which glutamine side
chains in protein substrates are deaminated via acyl-S-transaminase
intermediates, which are captured by Lys-e-NH2
groups to effect net transamidations (Scheme 15).[56,57]
3. Covalent Addition: The Supporting Cast
While the five chemical types of posttranslational modifications
discussed in Section 2 are abundant and wellcharacterized
in cells, there are many additional classes of
purposeful enzymatic modification of proteins that expand
the metabolic and signaling capacities of organisms.
3.1. Protein Hydroxylation
One additional category of protein posttranslational
oxidation is enzyme-mediated hydroxylation. Hydroxylations
occur at nonnucleophilic sites in aminoacyl side chains to
generate 3-OH-Pro, 4-OH-Pro, and 5-OH-Lys (Figure 10) in
collagen at Pro-Gly and Lys-Gly sites. These hydroxylations
are key modifications for proper maturation of collagen
fibers.[58]
Some of the 5-OH-Lys residues are then subsequently
tandemly glycosylated on the
newly introduced OH group to create
an O-disaccharide linkage. 4Hydroxyproline
modifications, about
tenfold more abundant than the
hydroxylations at C3 of Pro residues,
are involved in the triple helical
strands of collagen, with the 4-OH
pointing away from the helix.
A third side chain in which a CH2
group is converted into CH-OH is
Scheme 14. Oxidation of thiolate side chains of cysteine residues:
a) oxidation of dithiols to disulfides (for example via sulfenic acid
intermediates) and reversible reduction back to dithiols by glutathione
reductase action; b) oxidation of Cys-SÀ
side chain to S-nitrosyl-Cys by
nitric oxide (CNO).
Scheme 15. Nonoxidative cross-links introduced by transglutaminases; the amide bond in Gln
side chains is replaced by the e-NH of Lys to create Glu-e-Lys isopeptide cross-links.
Figure 10. Hydroxylated amino acid residues generated by posttranslational
FeII
-dependent monooxygenases: 3-OH-Pro, 4-OH-Pro, 5-OHLys,
3-OH-Asn.
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Asn (to 3-OH-Asn, Figure 10) in a small number of proteins,
including the transcription factor HIF (hypoxia inducible
factor).[59]
HIF transcription is initiated at low partial
pressures of O2 and induces the transcription of hundreds of
genes, including the gene that encodes erythropoietin to make
more red cells that can carry more O2 to hypoxic tissues.
The HIF-1a subunit in the HIFab heterodimer is posttranslationally
hydroxylated by hydroxylases, one specifically
acting at two Pro residues to create 4-OH-Pro[60]
and the other
acting at a particular Asn residue to generate 3-OH-Asn.[61]
These are determinative changes in the oxygen-sensing
cascade in mammalian tissues (Scheme 16).[62]
The lifetime
of the HIF-1a subunit in cells is controlled by proteolysis in
the ubiquitylation pathway, which involves chaperoned escort
to the proteasome, unfolding, and threading into the proteasome
chamber for proteolysis to limit peptides. The polyubiquitylation
of HIF-1a is carried out by a particular E3
ligase, the von Hippel-Lindau (VHL) protein.[63,64]
The
hydroxylation status of Pro402 and Pro564 in HIF-1a controls
the affinity for the VHL E3 ligase.[60]
At low pO2 in cells the
Pro hydroxylase is not saturated with its substrate O2 and has
low activity. At high pO2 the hydroxylase is active and
converts the two Pro residues into 4-OH-Pro residues. The
hydroxy side chain in HO-Pro564 provides about a 1000-fold
tighter binding of modified HIF-1a over unmodified HIF-1a
for the VHL ubiquityl ligase.[60]
This is the mechanism for
selective polyubiquitylation of the hydroxylated forms of
HIF-1a, which leads to its proteasome-mediated destruction
at high pO2 levels but its persistence at low pO2. Persistence
means a longer lifetime for the heterodimeric HIF-1ab and
the longer gene-transcriptional-activation response characteristic
of hypoxia.
The protein hydroxylases that catalyze the side-chain
hydroxylations noted in this section belong to the family of
non-heme FeII
monooxygenases that have two His and one
Asp side chains to provide three of the six coordination sites
to the FeII
(Scheme 17).[65]
Two additional coordination sites
are filled by cosubstrate a-ketoglutarate and the sixth by O2.
When both O2 and a-KG are bound, the organic diacid is
oxidatively decarboxylated to succinate. O2 is cleaved in such
a way that one atom ends up in succinate and the other is
coordinated to the iron as a high-valent FeIV
= O. This highvalent
oxoiron complex is a sufficiently powerful oxidant to
cleave the unactivated CÀH bonds at C3 and C4 of Pro
residues, C5 of Lys residues, and C3 of Asn residues to
generate transient CCH radicals and FeIII
-OH. OHC transfer
from the FeIII
-OH to the carbon-centered radical yields the
hydroxylated protein side chains. The polarity of these sidechain
hydroxylations is quite distinct from the bulk of the
other posttranslational modifications considered in this
Review. The amino acid side chains undergoing modification
are not electron rich or nucleophilic. Instead, the iron-based
modifying enzyme generates a powerful oxidant and leads to
homolytic cleavage of unactivated CÀH bonds with regio- and
stereospecificity.
Scheme 16. a) Hydroxylation of Pro and Asn residues in the HIF-1a
subunit; b) interaction of the HO-Pro564 residue of HIF with the E3
ligase that will catalyze polyubiquitylation of HIF.
Scheme 17. Mechanism of protein hydroxylation by the nonheme FeII
monooxygenases.
The active site iron(ii) is coordinated by three residues from within the
protein (two His, one Asp). Subsequent coordination by a-ketoglutarate and O2
results in the cleavage of dioxygen and formation of a high-valent FeIV=O species.
This iron complex cleaves the unactivated CÀH bond of the substrate, yielding
hydroxylated product and succinate.
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3.2. Protein Sulfur Transfer
Phosphoryl groups are not the only inorganic moieties
transferred to protein side chains. The SO3
À
group is also
transferred from phosphoadenosine phosphosulfate (PAPS)
to tyrosine side chains in proteins.[66]
PAPS is the biological
reagent for transfer of activated sulfuryl (SO3
À
) groups to
nucleophiles in both small molecules (e.g. oligosaccharides
such as heparin[67]
) as well as proteins and proteoglycans.
Enzymatic sulfation of four tyrosine residues occur in the
Golgi complex at the N-terminus of the CCR5 receptor
(Scheme 18) during its transit to the plasma membrane[68]
where the N-terminus is displayed at the extracellular surface.
This cluster of anionic Tyr-OSO3
À
modifications is important
for recognition by the CCR5 chemokine ligand.
Once sulfated molecules are internalized, the sulfate ester
bond is hydrolyzed enzymatically by sulfatases in the
secretory compartment, predominantly in lysosomes.[69–71]
The set of sulfatases known to degrade aryl sulfate ester
substrates are themselves in inactive proenzyme forms until
they undergo posttranslational activation. This involves
oxidative conversion of an active-site cysteine residue into
an aldehyde in the form of formylglycine (Fgly).[70]
This
conversion of a thiolate nucleophile into an electrophilic
carbonyl is followed by a hydration equilibrium to distribute
the Fgly between the aldehyde and the gem diol, the aldehyde
hydrate (Scheme 19). It is the hydrated form of the aldehyde
group in the formylglycine side chain that initiates covalent
attack on sulfated protein substrates bound in the sulfatase
active sites[72]
with resultant OÀSO3
À
bond cleavage.
3.3. Protein Modification by Bacterial Toxins
Bacteria that invade eukaryotic cells secrete a complement
of proteins into the host cell to neutralize its defense
mechanisms. Among these virulence factors are three types of
enzymes that act as posttranslational modification catalysts
for ADP ribosylation, glucosylation, and deamidation of host
target proteins.[3]
Scheme 18. Transfer of four SO3
À
groups from PAPS to the phenolate oxygen atoms of four side chains of Tyr residues at the N-terminal region of
the CCR5 receptor during its passage through the secretory compartments to the cell surface.
Scheme 19. Oxidative conversion of an active site Cys-SÀ
to the
aldehyde of formylGly (Fgly) converts inactive precursors of sulfatases
to active catalysts. Hydration of the Fgly side chain generates a gem
diol, which is the active nucleophile that attacks the Ar-OSO3
À
substrate on sulfur to initiate the O-SO3
À
bond cleavage.
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3.3.1. ADP Ribosylation
Several famous bacterial exotoxins, including the cholera
toxin, the diphtheria toxin, the pertussis toxin, and the
botulinum C3 toxin,[73]
are ADP ribosyltransferases. The
donor substrate of the ADP ribosyl moiety is the readily
available coenzyme NAD. The positively charged nicotinamide
group departs in a transition state involving the
ribaoxacarbenium ion of the transferring ADP ribosyl
group (Scheme 20a). This ion can be captured by robust
nucleophiles in protein cosubstrates in the active site of the
toxin, such as thiolate side chains of Cys, for example when
pertussis toxin modifies the a subunit of the inhibitory
GTPase Gi that regulates cyclic AMP production. The
transferring ribaoxacarbenium ion can also be captured by
weak nucleophiles such as the guanidino group of Arg in the a
subunits of the Gs GTPase by the cholera toxin. Another
example of the capture of the potent electrophilic form of the
ADP ribosyl moiety is the modification of the weakly
nucleophilic Asn41 in the Rho subfamily of small GTPases
by the C3 toxin from Clostridium botulinum, ultimately
leading to a net depolymerization of the actin mesh work in
the host cell.
A fourth example of bacterial toxin ADP ribosyltransferase
activity on a specific host protein with deleterious
consequence is the action of the diphtheria toxin on His715
in the protein synthesis factor eukaryotic elongation factor 2
(eEF-2). That histidine in mature eEF-2 has already undergone
a preparatory set of posttranslational modifications
involving the aminocarboxypropyl transfer from cosubstrate
S-adenosylmethionine (SAM), N,N,N-trimethylation by three
additional SAM molecules, and glutamine-mediated amidation
to convert His715 into a diphthamide residue. This is the
molecular form of eEF-2 that undergoes ADP ribosylation by
the diphtheria toxin (Scheme 20b) and rendered
inactive in its essential elongation functions,
bringing host protein synthesis to a halt in
the infected cell.[73]
3.3.2. Other Modification Activities of Bacterial
Toxins
The Ras and Rho GTPase families are also
targets of additional bacterial toxin enzymes
that inactivate the GTPases by chemical modifications
distinct from ADP ribosylation. One
is the lethal toxin protein from Clostridium
sordelli which also is a protein glycosyltransferase.
In this case it is not transfer of the ADP
ribosyl moiety from NAD but instead the
transfer of a glucosyl group from UDP-glucose
that is catalyzed by the protein toxin. The target
nucleophile is the b-OH of Thr35 in Ras. This Oglucosylation
blocks the catalytic activity of
Ras.[74,75]
A distinct type of posttranslational modification
strategy is deamidation of Gln61 in Rho
by the cytotoxic necrotizing protein from
pathogenic strains of E. coli.[76]
This Gln carboxamide
side chain is in the GTPase active site
and its hydrolysis to the g-COOÀ
of Glu61
disrupts the active-site machinery. Altogether,
the small GTPase families, because they act as
thermodynamic switches at so many cellular
intersections of signaling and metabolism, are
subjected to a diverse set of programmed
chemical modifications. The covalent modifications
alter localization and control function
both in normal maturation and in pathogen
interceptions.
3.4. Installation of Swinging-Arm Cofactors
Several enzymes central to primary metabolism
are nonfunctional until posttranslationScheme
20. ADP ribosylation of protein substrates by bacterial toxins: a) Cleavage of
NAD releases nicotinamide and generates a stabilized ribaoxacarbenium ion in the
active site. This can be captured by a range of cosubstrate nucleophiles, from the
robust Cys-SÀ
in pertussis toxin catalysis to the weak Asn and Arg side chains in
botulinum and cholera toxins. Water is a natural nucleophile and leads to NAD
glycohydrolase side reactions; b) diphtheria toxin catalyzes ADP ribosylation of a
modified histidine residue in the eukaryotic host cell protein synthesis elongation
factor eEF-2. The residue, His715 in the nascent eEF-2 undergoes five posttranslational
steps of transfer of the methionyl moiety from SAM, N,N,N-trimethylation, and
amidation to yield the diphthamide residue at position 715. This is the residue
targeted for ADP ribosylation on N3 of the imidazole ring by the diphtheria toxin.
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ally primed with prosthetic groups that provide key functional
groups to enable acyl- and carboxyl-transfer chemistry. The
acyl-transfer coenzymes are lipoic acid and phosphopantetheine.
The carboxyl-transfer prosthetic group is biotin
(Scheme 21). All three cofactors are covalently attached to
side chains of target apo proteins by dedicated activating/
loading enzymes. Biotin and lipoate can be activated as the
acyl AMP species and captured by the e-NH2 of lysine side
chains presented in folded 100-residue domains of the target
proteins to create the biotinylamide and lipoamide linkages.
The full reach of the Lys biotin and Lys lipoate chains is about
20 Š, leading to the historical connotation as swinging-arm
prosthetic groups that can visit different domains in multienzyme
complexes.[77]
In analogous logic the phosphopantetheinyl
moiety of CoASH as donor substrate can be captured
by the b-OH side chain of a specific Ser side chain in a 80–100residue
acyl-carrier protein domain, again to create a tethered
prosthetic group on a 20-Š pivot. In this case the chemical
bond is not an amide linkage but a phosphodiester.
The tethered biotinyl cofactor is used for the transfer of C1
groups in the form of CO2. In carboxylases that fix HCO3
À
into acetyl CoA and propionyl CoA, to yield malonyl and
methylmalonyl CoA products, there are multiple subsites or
distinct subunits with different chemical functions. In the
biotin carboxylase subunit, ATP is used as cosubstrate with
HCO3
À
to generate a transient carboxyphosphate mixed
anhydride that is captured by the biotinyl-Lys on the N1
ureido nitrogen atom to produce N1-carboxybiotinyl-Lysenzyme
(Scheme 22). This form of fixed CO2 is shuttled to the
active site where the C2 carbanion on the acetyl moiety of
acetyl CoA is generated and used to attack the N-carboxybiotinyl
tether. This action leads to CÀC bond formation as
acetyl CoA is carboxylated to malonyl CoA, one of the key
building blocks for fatty acid biosynthesis in cells.
The other two tethered coenzymes, lipoamide and pantetheinyl-phosphate,
are used to ferry substrate-derived acyl
groups between active sites of multidomain enzymatic
assembly lines.[77,78]
The lipoamide prosthetic group is found
in all a-keto acid dehydrogenase
complexes that carry out oxidative
decarboxylation, for example, of
pyruvate at the end of glycolysis
and of a-ketoglutarate in the citric
acid cycle. For example, in pyruvate
dehydrogenation the disulfide form
of the lipoamide cofactor is the
electron sink for ring-opening capture
by the C2 carbanion of hydroxyethylthiamine-PP
(Scheme 23). The
transferring two-carbon fragment
has been oxidized from the acetaldehyde
to the acetate oxidation state
and captured as an activated acetylS-lipoamide
thioester as the disulfide
link of oxidized lipoamide is
reduced. This is the prototypic
redox/energy-capture role for the
lipoamide prosthetic group. In the
second half-reaction, the acetyl-Slipoamide
arm moves to a separate
active site and docks next to a
CoASH molecule, thus allowing
acetyl transfer to the thiolate of
CoASÀ
, an isoenergetic transfer
that now releases the oxidized
acetyl moiety as the diffusible cellular
energy currency acetyl CoA.
A third example of cofactor
tethering, the pantetheinyl moiety
on acyl-carrier protein domains
(ACPs), also offers a terminal nucleophilic
thiolate for capture of acyl
groups. Its central role in primary
metabolism is in fatty acid biosynthesis
in which acyl chains are built
up by two carbon atoms at a time by
cycles of Claisen condensation followed
by redox tailoring to convert
Scheme 21. Coenzymes that are tethered in the active sites of enzymes to act as swinging-arm prosthetic
groups, carrying CO2 or acyl groups between active sites: biotin and the resultant biotinylamide-Lys; lipoate
and the resultant lipoamide-Lys; CoASH and the resultant pantetheinyl-OPO3-Ser linkages.
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elongated beta keto acyl-S-ACP
intermediate into the beta methylene
acyl-S-ACPs for the next
round of C2-unit elongation
(Scheme 24).[78]
The acyl groups
are proposed to visit ketosynthase,
ketoreductase, dehydratase,
and enoyl reductase active
sites sequentially while tethered
to the flexible phosphopantetheinyl
arm.
It would appear that Nature
has invented the coenzyme covalent
tethering strategy in these
three cases to fix small-molecule
acyl fragments or CO2 and ferry
them to distinct sites on multienzyme
complexes and assembly
lines. There are other examples in
which cofactors are tethered to
lysine side chains of proteins,
through hydrolyzable imine links
between the Lys-NH2 and an
aldehyde carbon atom in the
cofactor. This is true for the
aldehyde form of vitamin B6, pyridoxal
phosphate, in all pyridoxal
phosphate dependent enzymes,
and in the visual pigment proteins,
the rhodopsins, in which
vitamin A aldehyde (retinal) is
the aldehydic chromophore.
3.5. Posttranslational Carboxylation
of Glutamyl Residues for
Bidentate Calcium Binding
A family of proteins involved
in blood coagulation in mammals
undergoes posttranslational modifications
at sets of closely spaced
glutamate residues during passage
through secretory compartments
on their way to the extracellular
space. These modifications
involve fixation of CO2 to the gmethylene
carbon atoms of Glu
residues, thus creating malonyltype
side chains, known as gcarboxy
Glu (Gla). The Gla side
chains provide the possibility for
bidentate chelation of divalent
cations, of which interaction with
Ca2+
ions is most significant. The
proteins that undergo tandem gGlu
carboxylation include the
proenzyme forms of proteases
such as prothrombin, proFacScheme
23. The oxidized disulfide form of lipoamide
is the electron sink for capture by the
C2 carbanion of hydroxyethylthiamine pyrophosphate
(HE-TPP), resulting in release of the TPP
thiazole carbanion and generation of acetyl-Slipoamide.
The transferring two-carbon-atom
fragment has undergone oxidation, the lipoamide
disulfide has undergone reduction, and
energy has been captured in the acetyl thioester
linkage, which is maintained in subsequent
transfer of the acetyl moiety to CoASÀ
.
Scheme 22. N-carboxylation of biotinylamide by ATP and HCO3
À
in the biotin carboxylase active site
followed by ferrying of the tethered CO2 to the acetyl CoA carboxylation active site for CÀC bond
formation as malonyl CoA is produced.
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tor IX, and proFactor X.[79]
The carboxylation of 10–12
glutamate side chains to g-carboxy-Glu residues in a 40residue
stretch of the proenzyme forms of proteases creates a
local high density of bidentate chelators for Ca2+
ions
(Figure 11). The conformations of the Gla domains are
altered in the presence of Ca2+
and drive the association of
the proteases on platelet surfaces, leading to the formation of
protein complexes and the activation of neighboring proteases
to initiate and propagate blood coagulation cascades.[79]
Unlike the carboxylation reactions discussed in Section
3.4, which use tethered biotin as the CO2 carrier between
the active sites of the enzymes, the Gla-forming carboxylations
of glutamyl residue side chains do not use biotinyl amide
dependent enzymes. Instead, the naphthoquinone vitamin K,
in its only well-defined role in mammalian metabolism, is the
requisite cofactor for CO2 fixation in the Gla modifications.
In fact it is the dihydro-naphthoquinol form of vitamin K, that
is the active form for the vitamin K dependent protein
carboxylase. O2 is also a required cosubstrate and the
mechanism of the carboxylase (Scheme 25) is proposed to
involve formation of the K-OOH peroxy adduct. Cyclization
of this quinone hydroperoxide to the alkoxide anion of the
2,3-epoxide of vitamin K generates the strong base required
to abstract a proton from the glutamyl-g CH2 side chains. The
carbanion generated is required to attack CO2 and form the
new CÀC bond of the malonyl side chains of Gla residues in
the product.
4. Cataloguing the Posttranslational Modification
Consideration of major and minor categories of posttranslational
modifications above leads to at least hundreds of
thousands, perhaps millions, of possible molecular variants of
proteins in eukaryotic cells. The analytical problem is
immense, and the challenge to integrate across the subproteomes
and decipher connections for a systems biology
perspective even greater. Much of the contemporary effort
in PTM research involves the development of methodologies
to evaluate such inventories. Mass-spectrometric approaches
are dominant owing to issues such as femtomolar sensitivity of
detection and simultaneous identification of many peptide
fragments bearing a particular type of chemical modifica-
tion.[80]
Notable recent studies include the detection of
hundreds of pS and pT peptides from the phosphoproteome
of yeast[81]
and the isolation of His-tagged ubiquitylated
proteins of yeast. The latter method allows the identification
of more than 100 ubiquitylated proteins that contain polyubiquityl
chains connected through different Lys residues in
the ubiquitin monomers.[82]
At any moment in time, sampling of the proteome in a
given organism or cell provides only a snapshot of a highly
dynamic process, confounding the analytical problem and
ultimately arguing for time-resolved inventories. Heterogeneity
can arise in several ways. Because posttranslational
modification enzymes do not work off templates, the modScheme
24. The terminal thiolate of the phosphopantetheinyl prosthetic
group is the nucleophile that captures the starting acyl fragment
in fatty acid biosynthesis and serves as the platform on which the acyl
chain is elongated by two carbon atoms in each subsequent cycle. The
pantetheinyl arm carries the growing acyl chain between the active
sites of ketosynthase, ketoreductase, dehydratase, and enoyl reductase
in each cycle, converting the b-ketone into the four-electron-reduced
b-methylene oxidation state.
Figure 11. Twelve closely spaced Glu residues in the carboxylation
domain of the pro form of the coagulation protease factor IX are
modified to g-carboxy-Glu (Gla) residues and provide a high local
concentration of bidentate malonyl side chains for Ca2+
coordination.
The 3D fold of dodecaGla region complexed to eight calcium ions
(yellow spheres) emphasizes the divalent metal-ion-dependent structuring
of this region of the protein.
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ifications are stochastic and likely to be incomplete across a
target protein population. The fractional efficiency of modification
of a site, for example, N-glycosylation of a particular
Asn residue in an Asn-X-Ser sequence favorable for modification,
may depend on the amount of time the protein
region bearing that Asn is unfolded during passage across the
ER membrane. The accessibility of the carboxamido nitrogen
atom may be fleeting. The concentration of oligosaccharyl-Ntransferase
in the microenvironment may be limiting such
that the dwell time of protein substrate and enzyme catalyst is
too short to ensure complete modification at 100% of that site
in the protein population. In the multistep maturation of Nglycan
Man3(GlcNAc)2 core in glycoproteins there are up to a
dozen such substrate–catalyst encounters. A 90% yield at
each step would produce the kind of complex and heterogeneous
glycoprotein mixtures that are often detected. As noted
earlier, the prion protein has been shown to exist in some 52
glycoforms.[45]
5. Multiple and Tandem Posttranslational
Modification of Proteins
Posttranslational modifications often occur at
multiple sites or in tandem cascades that are crucial
for function. Thus, the Abl tyrosine kinase is found
phosphorylated at 11 different sites, (nine tyrosines,
one serine, one threonine) spread over the different
catalytic and regulatory domains of the protein[83]
(see Figure 2). In principle, 11! = 40420800 distinct
phosphorylated isoforms are possible just for this
one protein alone. Fractional occupancy is likely at
the distinct sites, making for a large nested array of different
phospho forms of this one protein.
Multiple lysine acetylations and methylations, one serine
phosphorylation, and one N-terminal ubiquitylation are
typical for the tails of the histone octamers in nucleosomes
so that modifications at 28 sites are possible as described in
Sections 2.2 and 2.3. (Surely not all of the 28! possibilities will
have been explored in Nature.) Kelleher and co-workers[84]
recently devised bioinformatic methods coupled with highresolution
mass spectrometry to predict, search for, and
identify particular histone variants. For example, they
detected one particular hexamodifed form of the N-terminus
of histone H3 with an [M+238] mass signature (Figure 12) to
address such a functional covalent diversity of protein
isoforms. (This is the equivalent of finding one needle in a
haystack. The systems approach to PTMs would be to assess
how many such needles altogether are in the “haystack” of
proteins.) The multiple modifications of histone tails are
Scheme 25. Proposed mechanism for the function of the dihydro form of vitamin K (KH2) as cosubstrate in the Glu to Gla posttranslational
protein carboxylations. KH2 reacts with cosubstrate O2 to produce a hydroperoxy vitamin K intermediate, which can proceed to the 2,3-epoxyvitamin
K alkoxide. The alkoxide is argued to be a strong enough base to abstract one of the Glu g-CH2 hydrogen atoms as a proton, transiently
generating the g-carbanion required to attack CO2 to produce the new CÀC bond in the Gla product residue.
Figure 12. Tandem enzymatic posttranslational tailoring of the N-terminal tail of histone H4
generates a heptamodified protein in which the N-terminal residue is acetylated,
lysines5,8,12,16 are acetylated, and lysine20 is N,N-dimethylated.
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presumed to be written and read by enzymatic machinery in
specific temporal patterns for selective recruitment of transcriptional
repressors and activator complexes.
Tandem posttranslational modifications are known in
many other proteins and comprise the multilayered informational
content driving the molecular logic of posttranslational
modifications. The consecutive four-step modification of the
C-termini of Ras proteins—1) S-prenylation, 2) S-palmitoylation
of neighboring cysteine residues, 3) specific endoproteolytic
cleavage to reveal one of the cysteines as the new Cterminus,
and 4) methylation of that new C-terminal carboxylate—comprise
the integrated maturation
process that moves modified Ras
to membranes to dock with its
upstream protein kinase partners
(Scheme 26).[35]
We have noted above
in Section 3.3 that the related RhoA
family of GTPases can undergo ADPribosylation,
O-glucosylation, and deamidation
of the active-site Gln. Adding
in this sequence of prenylation, proteolysis,
and C-terminal carboxymethylation
for Rho brings the total to seven
specific steps of posttranslational modification.
The mature RhoA can also be
cleaved by the YopT protease from
Yersinia pestis as that pathogen executes
part of its virulence program.[85]
The threshold nature of tandem
posttranslational modifications to provoke
a biological signal is illustrated
clearly in the successive addition of a
minimum of four ubiquitin 8-kDa tags to a target protein to
set off the cascade of events leading to proteolysis. A
polyubiquitin moiety of length at least four ubiquitin units
appears to be the threshold for recognition by protein
chaperones to send the tagged proteins to proteasomes.[27]
Finally, there is good evidence for competition between
posttranslational modifications, with opposing functional
consequences for the target proteins. Two such examples
involving competition between ubiquitylation and acetylation
are 1) SMAD7 protein in TGFb signal transduction path-
way[86]
and 2) five lysine side chains near the C-terminus of
the transcription factor p53. The Lys e-NH2 residues can be
acetylated or ubiquitylated (Scheme 27) and then extended to
polyubiquitin chains, leading to proteolytic removal of p53 or
SMADs. The acetylations block ubiquitylations and consequently
lengthen the lifetime of the proteins in cells.[13]
6. Reversible versus Irreversible Posttranslational
Modification
Depending on the biological purpose of a particular
covalent modification of a protein, reversibility may or may
not be an important parameter to
control. The prototype of reversible
modification is protein phosphorylation,
consistent with its evolution to
the dominant role in protein-based
signaling in eukaryotes. Of the five
major categories of PTMs noted in
Section 2 (phosphorylation, acylation,
glycosylation, thiol-disulfide chemistry,
and alkylation), all but alkylation
have dedicated enzymes, often large
enzyme families, that catalyze the
removal of the covalent modifications.
The enzymes that reverse phosphorylation,
acylation, and glycosylation are,
by and large, specific hydrolases,
whereas disulfide bonds are cleaved
by reductases. We note below that an
oxidative enzymatic route has now
been discovered for alkylation that
involves removal of N-methyl substituents
from the histone tails.
Scheme 26. Multistep modification of the Ras GTPase involves a) S-prenylation, b) endoproteolysis, and
c) C-terminal O-methylation.
Scheme 27. Competition between acetylation and ubiquitylation at the e-NH of Lys370,372,373,381,382
near the C terminus of the transcription factor p53.
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Protein kinases usually occur as low-activity “off” forms in
basal states in the absence of a specific stimulus. When a
signal is propagated and the kinases activated, often involving
covalent autophosphorylation of Thr residues to pT or TXY
sequons to doubly phosphorylated pTXpY loci in MAP
kinases, the activated protein kinases modify their partner
target proteins with much higher catalytic efficiency.[8]
The
cyclic AMP activated protein kinase A, for example, phosphorylates
over 100 proteins[87]
on Ser and Thr residues to
propagate signals.
To control the duration and intensity of phosphoprotein
signaling, the signals need to be turned off. Termination is
accomplished by hydrolytic removal of the PO3
2À
group by
phosphoprotein phosphatases (Scheme 28a). There are about
150 protein phosphatases encoded in the human genome,
including pS-, pT-selective phosphatases, pY phosphatases,
and dual-specificity (pS and pY) phosphatases.[88]
Of these,
107 are pY protein phosphatases.[89]
The much smaller
number of pS/pT phosphatases is balanced by the presence
of many regulatory subunits that control subcellular location
and substrate recognition for the pS/pT hydrolytic enzymes.
One practical consequence of PTM enzyme-mediated
reversibility is that the phosphoproteome content in any cell
at any given time represents the balance of activity of the 500
protein kinases/150 protein phosphatases towards their
diverse protein substrates, allowing an almost infinite
number of set points to a cell and true complexity for
phosphoproteomicists.[90,91]
Protein acetylation, protein ubiquitylation, and protein-Spalmitoylation
are three other classes of covalent modifications
that are readily reversed (Scheme 28b),
and so qualify for regulation and signaling
roles. Histone acetyltransferases (HATs) are
opposed functionally by a family of histone
deacetylases (HDACs).[92,93]
Some HDACs are
catalytic domains embedded in multimodular
proteins dynamically recruited to acetylated
and methylated lysine tails of histones. A
separate family of NAD-cleaving histone
deacetylases, the sirtuins, are involved in
gene silencing functions and couple energy
metabolism with transcriptional regula-
tion.[94,95]
Histone Lys-N-methylation, unlike Lysacetylation,
is not susceptible to enzymatic or
non-enzymatic hydrolytic reversal because
there is no obvious path for hydrolytic cleavage
of the N-alkyl bonds. Thus, in terms of
writing and rewriting the histone code, it has
been assumed that acetylation can be readily
erased, but not methylation. However, the
recent discovery of FAD-dependent methylLys
deaminases provides an oxidative route
for counteracting the action of histone meth-
yltransferases.[96]
Oxidative enzymatic conversion
of an N-methyl-Lys into the CH2=NH-Lys
product linkage (Scheme 29) now creates a
product imine labile to hydrolysis to yield the
unmodified H2N-Lys residue and 1 equivalent
of formaldehyde.
The dozens/hundreds of protein ubiquitin
ligases create ubiquityl-e-NH-Lys-protein isopeptide
bonds in tagged proteins. These linkages
are resistant to normal proteases but the
isopeptide bonds are cleaved by a family of
several dozen deubiquitylases (DUBs),[97,98]
presumably selective for subsets of ubiquitylated
proteins at different parts of the cell at
different times and rescuing those from proteasome
destruction.
Palmitoylated cysteine residues involve
covalent thioester linkages. They can hydrolyze
non-enzymatically, but this can be slow in
membranes and there are specific palmitoyl
Scheme 28. Reversibility of covalent modifications in proteins; a) Phosphoprotein phosphatases
oppose the effects of protein kinases; b) protein acetylations, S-palmitoylations,
and ubiquitylations are reversed by deacetylases, palmitoyl-S-protein thioesterases,
and deubiquitylases.
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protein thioesterases[11]
to control lifetime and consequent
localization of the protein at membrane interfaces.
In contrast to the above reversible categories of posttranslational
modifications are sets that are functionally
irreversible. Cys-S-prenylations are thioether linkages, and
in contrast to the Cys-S-palmitoyl thioester linkages are not
hydrolytically reversible. (Again, oxidative routes are known
but probably occur during prenylated protein degradation in
lysosomes.) Thus the enzymatic strategy for attaching the two
common lipid anchors for proteins, palmitoylation or prenylation,
provides reversible or irreversible modification of
proteins targeted to membranes. The C-carboxylation of Glu
residues to Gla residues is irreversible as is side-chain
hydroxylation, for example, of Pro and Asn residues on
HIF-1a in oxygen sensing.
The quintessentially irreversible posttranslational modification
is the second major category of covalent change to
proteins: the proteolytic cleavage of peptide bonds.
7. Controlled Proteolysis
The life cycle of every protein in intracellular and
extracellular compartments in an organism is controlled by
homeostatic functioning of proteases, which cleave the
covalent peptide backbones to release the constituent
amino acids back into the monomer pool. Large subsets of
proteins within eukaryotic cells may undergo consecutive
limited proteolytic clipping as part of the normal temporal
and spatial maturation process. These controlled proteolytic
cuts at specific peptide sequences within a given set of protein
substrates are effected by proteases that do not degrade but
rather are involved in protein-substrate maturation and
tailoring processes.
Essentially every protein that enters the endoplasmic
reticulum in eukaryotic cells undergoes cleavage of the Nterminal
25–30 amino acids, the signal sequence
that specified transit into the ER, by signal
peptidases.[99]
This first step in protein maturation
can be followed by action of proprotein
convertases later in the Golgi and trans Golgi
network[100,101]
of the secretory compartment. A
prototypical example is the cleavage of proinsulin
to insulin, whose two chains are connected by
disulfide bonds. The hormone cholecystokinin
undergoes six–eight cleavages and trimming
proteolytic maturations to convert the 115
residue initial translational product into the
eight-residue sulfated mature hormone.[102]
The maturation of the O-glycoprotein Notch
by limited proteolysis occurs in four spatially
and temporally distinct steps (Scheme 30):
1) signal peptide cleavage in the ER; 2) cleavage
by proprotein convertases into two subunits that
remain associated in the trans Golgi network on
the way to the cell surface; 3) removal of the
extracellular domain at the plasma membrane
by action of sheddase-type proteolytic activity,
triggered upon engagement of protein ligands;
4) regulated intramembrane proteolysis (RIP) of the truncated
Notch as the fourth maturation event.[99,103,104]
The
cytoplasmic stub of Notch, now with only a few remaining
Scheme 29. Reversal of lysine e-N-methylation occurs by oxidation (not hydrolysis) by action of
a flavoprotein to generate the hydrolytically labile imine product.
Scheme 30. Maturation of the Notch protein by four sequential proteolytic
cleavages at specific sites and specific places in a eukaryotic cell:
a) Cleavage of the N-terminal signal peptide upon secretion into the
endoplasmic reticulum (not shown in figure); b) hydrolytic clip by
proprotein convertases during passage through the trans Golgi network;
c) hydrolytic clip by a protein sheddase at the cell surface;
d) regulated intermembrane proteolysis releases the cytoplasmic stub
of Notch to go to the nucleus and act as transcription factor.
Reproduced with permission from reference [121].
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amino acids in the vesicle membranes, can then partition back
to the cytoplasm, enter the nucleus, associate with partner
proteins, and act as a selective transcriptional activator. The
sequence of four limited proteolytic steps, separated in time
and space, control the location and activity of Notch. It can be
directed through the ER and the Golgi complex to the cell
surface, function as a cell surface receptor, and then be
liberated back into the cytoplasm as a transcriptionally
activating fragment that can target the nucleus. Doubtless
there are many other examples of multistep action of
proteases to direct the location and function of client proteins
and control their lifetimes in cells.
A variant of controlled proteolysis occurs when a short Cterminal
peptide is excised from protein substrates by
protease-like catalysts that generate transient covalent proteinyl-S-Cys-enzyme
intermediates (Scheme 31). In the coupling
of GPI lipid anchors to eukaryotic proteins, the transferring
protein acyl group is specifically captured by an
ethanolamine group in the cosubstrate GPI anchor,[105]
a net
switching of the C-terminal peptide of the initial translation
product by the GPI anchor. In the action of bacterial
sortases,[106]
the incoming amine nucleophile that captures
the staphylococcal proteins to be displayed as antigens at the
bacterial cell surface is a Gly or Ala-NH2 terminus from crossbridges
in the peptidoglycan layer of the cell walls. Both of
these C-terminal modifications are net transamidations, in
which the incoming nucleophile is an amine to give an
aminolytic product rather than water in hydrolysis.
8. Autocleavage and Peptide-Bond Rearrangement
There is a set of autocatalytic processes that leads folded
proteins to catalyze rearrangements of the backbone connectivity
at one or more specific peptide bonds in the folded
proteins. Most often these rearrangements lead to cleavage of
Scheme 31. Protease-like catalysts use active-site Cys nucleophiles to transfer protein-substrate-derived acyl fragments to nonprotein amine
acceptors via acyl-S-Cys enzyme intermediates: a) GPI anchor attachment through enzymatic transamidation; b) cross-linking and display of
proteins to the peptidoglycan cross-bridges in Staphylococcus aureus cell wall assembly by transamidation by the enzyme sortase.
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those specific peptide bonds. The
fate of the acyl fragment of the
autocleaving peptide can vary
depending on the capturing
nucleophile. Water as cosubstrate
results in hydrolysis. An
alcohol as cosubstrate generates
an ester product, as in the capture
of the N-terminal fragment
of Hedgehog protein by cholesterol.
Capture by an amine,
when the amine is downstream
in the same protein, is the
essence of protein splicing,
although the amine capture is
indirect as we shall note. A
common type of intermediate is
observed for all three such capture
processes.
A second major variant of
autocatalytic peptide backbone
rearrangements occurs in two
contexts (Section 9). One is the
autoconversion of a tripeptide
moiety into the highly conjugated
aromatic fluorophore of
the green fluorescent proteins
and its relatives. The second is a
related rearrangement of a tripeptide
in the active site of the
pro forms of phenylalanine and
histidine deaminases to create
the imidazolone cofactors that
serve as essential electron sinks
in deaminase catalysis.
8.1. Autocleavage of Specific
Peptide Bonds in Proteins
During Precursor Activation
A relatively small subset of
folded proteins have the capacity
to convert themselves from a
single-chain inactive precursor
form into an active two-chain
form by hydrolysis of a specific
peptide bond in the precursor
(Scheme 32). Three examples of
variants of autoproteolytic activation
are: 1) cleavage of the
precursor single chain forms of
the b-subunits of proteasomes; 2) cleavage and uncovering of
an N-terminal pyruvamide group in aspartate a-decarboxylase;
3) cleavage and activation of Hedgehog family proteins
resulting in alcoholysis of the peptide bond by cholesterol. All
three enzymes use the side chain of a Ser, Thr, or Cys residue
to attack the immediately adjacent upstream peptide carbonyl
to generate a five-membered tetrahedral adduct
(Scheme 33). If that adduct is protonated on nitrogen and
reopens with cleavage of the CÀN bond, then the peptide
bond has been cleaved. The two parts of the protein chain are
still held together, by an O-ester (Ser, Thr) or an S-thioester
(Cys) linkage. This is the common thread of all three
examples. The O-ester or S-thioester linkage is now labile
to capture by even relatively weak nucleophiles such as water,
Scheme 32. Autoproteolysis at specific peptide linkages to create the active forms of enzymes in the
fragmented protein products; a) autocleavage of the precursor forms of the b subunits of proteasomes
to liberate the catalytic Thr nucleophile; b) autocleavage of the precursor form of aspartate decarboxylase
generates the N-terminal pyruvamide electron sink; c) autocleavage of the Hedgehog precursor
and capture by the 3-OH of cholesterol generates the active N-terminal fragment as a membranetethered
cholesterol ester.
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the alcohol group of cholesterol, or downstream Ser and Cys
side chains.
The proteasome is organized in an a7b7b7a7 four-layered
cylinder.[107]
In yeast, three of the seven b subunits in each
heptad ring are active, arising by autocleavage of the
precursor b chains at Gly75–Thr76, releasing the Thr as the
new N-terminus of the cleaved b subunit.[108–110]
Cleavage
occurs when the side chain of Thr76 attacks the adjacent
peptide carbonyl to intiate cleavage. The liberated amino
group of this new Thr1 is the nucleophile in catalysis as protein
substrates are cleaved, explaining why the precursor is
inactive.[111]
The proteasomal b subunit is one of several
proenzymes that autocleave and liberate the N-terminal
nucleophile as the catalytic residue.[109]
A related class of precursor forms of enzymes uses the
side chain CH2OH of a specific serine residue to initiate
cleavage in the folded precursor on the adjacent upstream
peptide carbonyl. As noted above, the tetrahedral adduct
decomposes to an O-ester, cleaving the original CÀN linkage
of the peptide bond. Elimination of the RO group from this
ester yields an N-terminal dehydroalanyl residue at the
downstream chain of the two-chain product (Scheme 34).[109]
The N-terminal dehydroalanyl moiety hydrates and ketonizes
to accumulate as a pyruvamide group at the N terminus of the
active enzyme. This directed autoproteolytic process has
uncovered a ketone moiety, an N-terminal electrophile rather
than the N-terminal nucleophile above, which acts as an
active-site electron sink for the substrate aspartate. Decarboxylation
of the aspartyl imino enzyme followed by hydrolysis
of the bound product imine yields b-alanine, a required
intermediate in CoASH biosynthesis.[112]
The third variant of precursor protein autoproteolysis is
represented by the maturation of the protein Hedgehog that
signals at plasma membranes of eukaryotic cells after twodimensional
diffusion in the plane of the membrane. The
protein is tethered to the membrane by a C-terminal covalent
lipid anchor, in this case by esterification of a cholesterol
moiety with the C-terminal carboxylate.[113]
This arises from
the autocleavage intermediate common to the above two
cases in which rearrangement of a peptide bond in the
precursor to an oxoester occurs by an identical mechanism.
Now, rather than capture of the acyl fragment of the precursor
protein by water for net hydrolysis, there is a specific binding
site for cholesterol. Its 3’-OH is the kinetically competent
nucleophile, yielding the peptidyl cholesterol ester as the
cholesterolysis product that is biologically active (see
Scheme 32).
These three variations of folded protein-precursor autocleavage
in Scheme 32 yield either a) two normal peptide
fragments, b) a downstream fragment with an N-terminal
pyruvamide, or c) an upstream fragment with a C-terminal
cholesterol ester. This shows the versatility of this autocatalyzed
peptide bond route by selective control of the fate of the
common rearranged ester intermediate. In all three cases the
peptide bond fragmentations are irreversible.
8.2. Autocleavage and Religation: Protein Splicing
In the final variant of autocleavage of protein precursors
the ester intermediate (or thioester intermediate when a Cys
thiolate attacks the adjacent upstream peptide bond) is
captured by an intramolecular nucleophile rather than an
external one such as HOH or ROH. The internal nucleophile
is a side chain Ser, Thr, or Cys in a folded downstream domain
of the precursor protein. This is the essence of protein
splicing, autocleavage, and peptide-bond religation, practiced
by over a hundred bacterial and yeast proteins,[109,114]
including
DNA polymerases and ATPases. The first peptide bond is
Scheme 33. Peptide-bond autocleavage mechanism proceeds via a) a
tetrahedral cyclic adduct arising from attack of a nucleophilic side
chain on the immediate upstream peptide bond; resolution of the
adduct proceeds through CÀN bond cleavage, breaking the original
peptide bond and generating an oxo/thioester connectivity; b) the
ester can be captured by a range of nucleophiles, including water.
Scheme 34. Peptide autocleavage at an X-Ser linkage can be followed
by elimination of the Ser-b-OH and tautomerization to an N-terminal
pyruvamide group. This is an electron sink that can engage in imine
formation with amino acid substrates to facilitate kinetically accessible
substrate carbanions, in this case for net decarboxylation of Asp.
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cleaved at the junction of upstream extein and intein. The
internal nucleophile capturing the extein1-intein-O-ester
intermediate is at the boundary of the same intein and
extein2 (Scheme 35). The new intermediate has a lariat
structure. Excision of the intein is completed with participation
of an Asn side chain at the downstream boundary of the
intein. This yields a second ester (thioester) intermediate as
the lariat is resolved. Re-formation of the peptide bond
joining extein1–extein2 is driven by thermodynamically
favored acyl-O or acyl-S to N shifts and completes an inframe
ligation.
This is ancient protein biochemistry in an evolutionary
sense (found in archaeal proteins) and may be the harbinger
of the other autocleavage reactions noted in the above
section. The net reversibility of such peptide-bond cleavage is
the remarkable outcome and is controlled by structural
features in the folded intermediates to keep water from being
a competent, interfering nucleophile to the intramolecular
acyl transfers within the self-splicing proteins. To the extent
that this is ancient protein chemistry, it emphasizes that acyl
oxoester and acyl thioester intermediates were primordial
species in protein reactions; they are still intermediates in
many posttranslational reactions in contemporary protein
maturations. Furthermore, protein splicing has many practical
applications in modifications of recombinant proteins with
synthetic peptides by protein ligation.[109,115]
9. Peptide Bond Rearrangement without
Autocleavage
9.1. Fluorophore Formation in Green Fluorescent Protein
An additional spectacular class of autocatalyzed PTM
rearrangement of the peptide backbone in folded proteins is
the maturation of chromoproteins of the green fluorescent
protein (GFP) and red fluorescent protein families (Scheme
36a).[116]
The precursor protein folds into a b cyclinder
structure and the native conformation is required for the
subsequent chromophore generation. A tripepetide loop of
Ser65Tyr66Gly67 in the folded colorless GFP precursor is
sterically compressed, populating a conformer that allows
attack of the Gly67 amide N-H on the adjacent peptide
carbonyl to generate a five-membered tetrahedral adduct
reminiscent of the initial steps in the rearrangements
described in Section 8. This adduct is dehydrated, and the
resultant stable cyclic species is slowly autoxidized to create a
double bond in conjugation with the phenol ring of Tyr66. This
oxidative last step generates the chromophore with absorption
maximum at 506 nm and the green fluorescence useful to
the producing coelenterate for energy-harvesting functions. A
variant with QYG in the starting tripeptide is found in related
coelenterates and yields the DsRed fluorophore after rearrangement
and oxidative maturation (Scheme 36b).[117]
FluoScheme
35. Autocleavage, intein excision, and peptide religation during protein splicing: thioester formation during peptide bond autocleavage,
followed by transthiolation to the cysteine at the intein–extein2 boundary creates a lariat intermediate, that resolves by participation of the Asn at
the C terminus of the intein. The resultant thioester reforms the extein1–extein2 peptide bond by acyl S to N shift.
C. T. Walsh et al.Reviews
7368 www.angewandte.org  2005 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim Angew. Chem. Int. Ed. 2005, 44, 7342 – 7372
rophores with altered colors, including cyan and gold, have
been generated by protein engineering and evolution
approaches, allowing many fluorescence-energy-transfer
studies between pairs of variant GFPs.
9.2. The Methyleneimidazolone Chromophore
Related rearrangements of a tripeptide sequon in a loop
of the folded proenzyme forms of phenylalanine and histidine
deaminases produce a related cyclic imidazolone group that
then functions as the activesite
electrophile for substrate
amine deamination (Scheme
37a).[118,119]
In the Pseudomonas
histidine deaminase,
Ala142Ser143Gly144 in a loop
region autoconverts to a compact
heterocycle in which the
Gly144 amide nitrogen is proposed
to attack the carbonyl of
Ala142. The cyclic tetrahedral
adduct can lose water and
form the C=N of the imidazolone.
Loss of the -OH from the
Ser143 side chain forms the new
double bond in conjugation to
produce the 4-methylene-5imidazolone
(MIO) prosthetic
group. This set of transformations
fashions an electrophilic
heterocyclic cofactor from the
tripeptide loop to generate the
active enzyme. The MIO heterocycle
in the mature, active
enzyme could be attacked by
the substrate amino group. An
elimination reaction initiated
by b-H removal would then
generate urocanate and the
amino moiety still attached to
the cofactor (Scheme 37b).
Release of NH3 regenerates
the starting MIO cofactor for
the next catalytic cycle.[120]
10. Conclusions
This summary of posttranslational
modifications of proteins
has not attempted
exhaustive coverage of the
more than 200 known covalent
modifications (see reference
[3] for more complete
coverage). Rather, emphasis
has been on how the addition
of chemical groups from
common cofactors and coenzymes to side chains of 15 of
the 20 amino acids found in proteins expands the proteome
structurally and functionally. The dramatic enhancement of
the capabilities of the limited scaffold of genetically encoded
protein backbones creates new functional capacities. The
modified proteins now have expanded opportunities for
catalysis, initiation and termination of signal cascades, integration
of information at many metabolic intersections, and
alteration of cellular addresses. The posttranslational diversification
of the proteome illuminates the underlying molecular
logic for epigenetic acquisition of new protein functions.
Scheme 36. a) Autoconversion of Ser65Tyr66Gly67 in the precursor of green fluorescent protein (GFP) to the
green fluorescent form: formation of a cyclic tetrahedral adduct followed by dehydration generates the
heterocycle that is not yet fluorescent. Extension of the conjugated system is a slow, oxygen-dependent
oxidation that brings the Tyr chromophore into conjugation and completes fluorophore formation. b) A
related protein with a Gln66Tyr67Gly68 tripeptide instead of the SerTyrGly sequence is the fluorophore in the
DsRed protein from coral. It proceeds through a green intermediate on the way to the final red form.
Protein Modification
Angewandte
Chemie
7369Angew. Chem. Int. Ed. 2005, 44, 7342 – 7372  2005 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim www.angewandte.org
The controlled cleavage of specific peptide bonds in
particular protein substrates, especially those transiting
eukaryotic secretory pathways, shows explicitly that proteases
need not be blunt instruments in cellular control inventories.
The life cycles of proteins secreted to the cell surface, engaged
by ligands, and then retrieved to the nucleus by consecutive
rounds of limited proteolysis show ingenious and sophisticated
use of the molecular logic of proteolytic trimming of
precursors.
Proteins can clearly help themselves for posttranslational
creation of new functional groups. Folded protein precursor
forms exhibit remarkable properties for autocatalytic conversion
of peptide loops into cyclic derivatives for catalysis or
light-energy harvesting. Controlled autocatalytic cleavages,
presumably vestigial capacities of proteins during evolution,
set up N-terminal nucleophiles, N-terminal electrophiles
(pyruvamides), and C-terminal cholesterol esters by routing
common intermediates down different decomposition pathways.
The apotheosis of these autocleavages are the error-free
religations in protein splicing reactions
in which inteins are excised and exteins
self-splice in frame to convert inactive
precursors into mature proteins. Splicing
may have been an important route
to shuffling domains within proteins as
a part of multidomain protein evolution
pathways.
A given protein can be posttranslationally
modified at many residues
with the same group, for example,
eleven phosphorylations of Abl, five
acetylations at the C terminus of p53.
Or a protein can be subjected to
tandem modifications by several
kinds of covalently introduced groups,
as exemplified by the two lipidations of
Ras followed by regiospecific proteolysis
and C-terminal O-methylation, or
the multiple modifications of Rho by
bacterial protein toxins that disrupt
eukaryotic cell cytoskeletal apparatus.
The coordinated orchestration of acetylations,
methylations, phosphorylations
and ubiquitylations of histone
tails on nucleosomes give insight into
the finely tuned molecular logic of
protein posttranslational modifications
for gene-expression control.
With the advent of many variations
of high-resolution mass spectrometry it
is possible to detect and localize covalent
changes in proteins beyond the
genetically encoded sequence. Cataloguing
the protein variants in various
subproteomes (such as the phosphoproteome,
the ubiquitylated proteome,
the molecular variants of histone modifications
in nucleosomes of differing
transcriptional activity) will continue
to be important parameters to allow a full description of the
protein composition of proteomes. This information will be a
necessary preamble to understanding how the diverse molecular
forms of proteins carry out their integrated functions.
We thank M. Fischbach for assistance with the production of
the frontispiece.
Received: March 21, 2005
Published online: November 3, 2005
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