Structural Studies of Human 5’-Nucleotidases Karin Walldén

Structural Studies of Human
5’-Nucleotidases
Karin Walldén
Doctoral thesis at Stockholm University
Department of Biochemistry and Biophysics
© Karin Walldén, Stockholm 2008
ISBN 978-91-7155-718-6
Printed in Sweden by Universitetsservice AB, Stockholm 2008
Distributor: Department of Biochemistry and Biophysics,
Stockholm University
Papers I-III are reprinted with permission from the publisher.
Abstract
5’-Nucleotidases (5’NTs) are catabolic enzymes of the nucleotide metabolism. They catalyze dephosphorylation of deoxyribo- and ribonucleoside
monophosphates and constitute an important control point in the regulation
of intracellular nucleotide pools for the maintenance of correct DNA and
RNA synthesis.
By removing the α-phosphate group from a nucleotide, the 5’NTs release
the nucleoside to pass the plasma membrane by facilitated diffusion. Depending on the cellular need for nucleotides, the nucleosides can either exit
the cell for reuse elsewhere or be imported and subsequently phosphorylated
by nucleoside and nucleotide kinases.
The knowledge of how nucleotides are metabolized has been used for rational design of nucleoside analogues that are used in treatment of cancer
and viral diseases. These drugs are phosphorylated within the cell to become
active. Their dephosphorylation by 5’NTs might be one of the mechanisms
behind the resistance experienced by patients towards such drugs.
This thesis describes structure-function studies on four of the seven
known human 5’-NTs. The focus of the work is on the substrate specificity
and regulation of these enzymes. Inactive variants of the mitochondrial and
cytosolic deoxynucleotidases and the cytosolic 5’-nucleotidase II were used
to characterize the structural basis for their substrate specificity in high detail.
Based on structures of the apoprotein and activator/activator+substrate
complexes of cytosolic 5’-nucleotidase II, a mechanism for the allosteric
activation of this enzyme was presented. In this mechanism, the activator
induces a conformational change that involves conserved residues of the
active site. The conformational change drastically increases the enzyme affinity for the phosphate moiety of the substrate.
List of publications
I
Wallden K., Ruzzenente B., Rinaldo-Matthis A., Bianchi
V. and Nordlund P. (2005) Structural basis for substrate
specificity of the human mitochondrial deoxyribonucleotidase, Structure 13, 1081-1088.
II
Wallden K., Rinaldo-Matthis A., Ruzzenente B., Rampazzo C., Bianchi V. and Nordlund P. (2007) Crystal structures of human and murine deoxyribonucleotidases: Insights into recognition of substrates and nucleotide analogues, Biochemistry 46, 13809-13818.
III
Wallden K., Stenmark P., Nyman T., Flodin S., Graslund
S., Loppnau P., Bianchi V. and Nordlund P. (2007) Crystal
structure of human cytosolic 5 '-nucleotidase II - Insights
into allosteric regulation and substrate recognition, Journal
of Biological Chemistry 282, 17828-17836.
IV
Wallden K. and Nordlund P. Mechanism for allosteric activation and substrate recognition of human cytosolic 5'nucleotidase II, Manuscript
Additional publication
Kosinska U., Wallden K., Flodin S., Nyman T., Stenmark P., Eklund
H. and Nordlund P. Structure of human uridine cytidine kinase 1 in
ligand free and ADP bound states, Manuscript.
Contents
Introduction .....................................................................................................1
Nucleotide Metabolism .................................................................................................... 1
De novo pathway ....................................................................................................... 3
Salvage pathway........................................................................................................ 4
Regulation of nucleotide metabolism .............................................................................. 4
Types of regulatory mechanisms............................................................................... 4
Regulation of de novo synthesis................................................................................ 5
Regulation of the salvage pathway............................................................................ 6
Cell cycle dependence............................................................................................... 6
5’-Nucleotidases (5’NTs) ................................................................................................. 7
Overview .................................................................................................................... 7
Cytosolic 5’(3’)-deoxynucleotidase (cdN) .................................................................. 9
Mitochondrial 5’(3’)-deoxynucleotidase (mdN) ........................................................ 10
Cytosolic 5’-nucleotidase IA (cN-IA) ........................................................................ 10
Cytosolic 5’-nucleotidase IB (cN-IB) ........................................................................ 11
Cytosolic 5’-nucleotidase II (cN-II) ........................................................................... 11
Cytosolic 5’-nucleotidase III (cN-III) ......................................................................... 12
Ecto-5’-nucleotidase (eN) ........................................................................................ 13
Nucleotide pools and disease ....................................................................................... 13
Nucleotide pools ............................................................................................................ 14
Nucleoside/nucleotide transporters............................................................................... 16
Anti-cancer/viral nucleoside analogues......................................................................... 17
Evolution and catalysis of intracellular 5’NTs................................................................ 20
Overall structure of intracellular 5’NTs..................................................................... 20
Catalytic mechanism................................................................................................ 23
Catalytic mechanism of eN ...................................................................................... 25
Present Investigation.....................................................................................27
Aim................................................................................................................................. 27
Strategy ......................................................................................................................... 27
Fluorometallic complexes as models for enzyme intermediates ............................. 28
Substitutions to trap substrate in active site ............................................................ 28
Structural basis for substrate specificity of mdN and cdN (Paper I and II) ................... 30
Pyrimidine base specificity....................................................................................... 32
Deoxy/ribo specificity ............................................................................................... 33
2’-,3’- and 5’-phosphate specificity .......................................................................... 34
Purine/pyrimidine specificity..................................................................................... 35
Nucleoside analogue recognition in mdN ................................................................ 36
Implications for the catalytic mechanism ................................................................. 38
Structure of human cN-II (Paper III and IV)................................................................... 39
Characterization of 2 effector sites .......................................................................... 39
Activator recognition in effector site 1...................................................................... 40
Mechanism for activation of cN-II ............................................................................ 42
Covalent modification on Asn52 .............................................................................. 45
Substrate recognition in cN-II................................................................................... 47
Structure of human cN-III .............................................................................................. 49
Mapping of cN-III deficiency substitutions ............................................................... 49
Phosphotransferase activity of cN-II and cN-III ....................................................... 53
Future perspectives.......................................................................................54
Acknowledgments .........................................................................................55
References....................................................................................................56
Abbreviations
Enzymes
ADA
AK/ UCK
ANC
APRT
cdN
cN-IA
cN-IB
cN-II
cN-III
CNT
DNC
dCK/ dGK
eN
ENT
HAD
HGPRT
mdN
(d)NK
NMPK/NDPK
5’NT
β−PGM
PNP
PSP
RNR
SERCA
TK1/TK2
TP/UP
Analogues
NA (-MP)
d4T
AZT
BVdU
FdU
AraT
ddC
Adenosine deaminase
Adenosine kinase/uridine cytidine kinase
Adenine nucleotide carrier
Adenosine phosphoribosyl transferase
Cytosolic 5’(3’)-deoxynucleotidase
Cytosolic 5’-nucleotidase IA
Cytosolic 5’-nucleotidase IB
Cytosolic 5’-nucleotidase II
Cytosolic 5’-nucleotidase III
Concentrative nucleoside transporter
Deoxynucleotide carrier
Deoxycytidine kinase/deoxyguanosine kinase
Ecto 5’-nucleotidase
Equilibrative nucleoside transporter
Haloacid dehalogenase
Hypoxanthine guanosine phosphoribosyl transferase
Mitochondrial 5’(3’)-deoxynucleotidase
(Deoxy)nucleoside kinase
Nucleoside mono-/diphosphate kinase
5’-Nucleotidase
β-Phosphoglucomutase
Purine nucleoside phosphorylase
Phosphoserine phosphatase
Ribonucleotide reductase
Sarcoplasmic Ca2+ ATPase
Thymidine kinase 1/ thymidine kinase 2
Thymidine/uridine phosphorylase
Nucleoside analogue (5’-monophosphate)
2’,3’-didehydro-2’,3’-dideoxythymidine
3’-Azidothymidine
E)-5-(2-bromovinyl)-2’-deoxyuridine
5-fluoro-2’-deoxyuridine
1-β-D –arabinosylthymine
2’,3’-dideoxycytidine
dFdC
araC
3TC
CdA
araG
DPB-T
PMcP-U
Diseases
AIRP
AML
MDS
MNGIE
Other
ApnA
GPI
Pi
2’,2’-difluoro-2’-deoxycytidine (Gemcitabine)
1-β-D –arabinofuranosylcytosine
2’-deoxy-3’-thiacytidine
2-chloro-2’-deoxyadenosine (cladribine)
9-β-D –arabinosylguanine
(s)—1-[2’-deoxy-3’,5’-O-(1-phosphono) benzylidene-bD-threo-pento-furanosyl]-thymine
(+)-1-trans-(2-Phosphonomethoxycyclopentyl)uracil
autoimmune infertility protein
acute myeloid leukemia
Mitochondrial DNA depletion syndrome
Mitochondrial neurogastrointestinal encephalomyopathy
Diadenosine 5’,5’-polyphosphate
glycosyl phosphatidylinositol
Inorganic phosphate
Amino acids
Alanine
Ala
Arginine
Arg
Asparagine Asn
Aspartic acid Asp
Cysteine
Cys
Glutamic acid Glu
Glutamine
Gln
Glycine
Gly
Histidine
His
Isoleucine
Ile
Leucine
Leu
Lysine
Lys
Methionine Met
Phenylalanine Phe
Proline
Pro
Serine
Ser
Threonine
Thr
Tryptophan Trp
Tyrosine
Tyr
Valine
Val
A
R
N
D
C
E
Q
G
H
I
L
K
M
F
P
S
T
W
Y
V
Introduction
Living organisms share the capacity to replicate themselves. To succeed with
this and to survive, they must take up energy from their surroundings and
converge this energy to synthesize organic material. This is mediated by
enzymes, which are the catalysts of life.
Humans contain thousands of different enzymes that catalyze specific
chemical reactions. Some enzymes degrade components of food and channel
the energy released by this process into chemical energy in the form of for
instance the nucleotide ATP. Other enzymes can then use ATP as energy
source for synthesis of various components of the cell.
The degradation (catabolism) and biosynthesis (anabolism) of an organic
compound usually involve several sequential reactions catalyzed by different
enzymes. These form together a “pathway”, in which one reactant (substrate)
enters and a product from the first enzyme will become a substrate for the
second enzyme and so forth.
Most such pathways are regulated, so that the amount of product is finetuned to the need of the cell. For instance, when exercising you activate
catabolic pathways that release sufficient amount of chemical energy needed
for the exercise. Enzymes can be regulated in several ways, in which either
the amount, localization or catalytic efficiency of the enzyme is changed.
The focus of this thesis is on a family of enzymes called 5’-nucleotidases
(5’NTs), which are catabolic enzymes that participate in the metabolism of
nucleotides.
Nucleotide Metabolism
Nucleotides (Fig. 1) play several important roles in all cells. ATP and to a
lower extent GTP are essential chemical energy carriers that transfer energy
liberated by catabolism to anabolic processes. Nucleotides are also used as
building blocks in DNA and RNA that carry the genetic information of
which proteins that can be produced by the cell. Nucleotides are also elements in cofactors, such as NAD, FAD and coenzyme A, and activated biosynthetic intermediates such as UDP-glucose (1).
1
Fig. 1. Composition of nucleotides. (a) The base: Adenine, guanine and hypoxanthine are modifications of purine, and uracil, thymine and cytosine are modifications
of pyrimidine. (b) The sugar ring: β−D-Ribose and β−D-deoxyribose. (c) Nucleoside: The four deoxynucleosides. d) Nucleotide: (Deoxy)nucleoside 5’-mono-, dior triphosphates, (d)NMPs, (d)NDPs and (d)NTPs.
Nucleotides are synthesized either de novo from low molecular mass precursors or from recycled nucleosides and bases. These two pathways are
called the de novo pathway and the salvage pathway, respectively (Fig. 2).
The 5’NTs are active in the salvage pathway.
Fig. 2. De novo pathway and salvage pathway.
2
De novo pathway
Nucleotides are composed of a nitrogen-rich base, a sugar ring and phosphate group(s) (Fig.1). The base is either a modified purine or a modified
pyrimidine that are synthesized onto a ribose 5’-phosphate by multi-step
enzymatic pathways from amino acids, CO2 and NH3 (1).
The de novo synthesis of pyrimidine nucleotides goes through the formation of uridine monophosphate (UMP) that is further converted to all other
pyrimidine nucleotides. The corresponding synthesis of purine nucleotides
leads to inosine monophosphate (IMP), which is the precursor to all other
purine nucleotides (Fig. 3).
Fig. 3. De novo synthesis of nucleotides goes through UMP and IMP.
Fig. 4 shows a scheme of the de novo pathway going from NDPs to
dNTPs for synthesis of DNA. Nucleoside monophosphate kinases (NMPKs)
phosphorylate the NMPs to NDPs, which are then reduced to dNDPs by a
single enzyme, ribonucleotide reductase (RNR) (2-4). Then nucleoside diphosphate kinases (NDPKs) phosphorylate dNDPs into dNTPs. An exception is the synthesis of dTTP, which is formed from dCDP through several
enzymatic steps (Fig. 4).
Fig. 4. De novo pathway: (1) RNR, (2) NDPKs, (3) dCMP deaminase, (4) Thymidylate synthase and (5) NMPKs
3
Salvage pathway
Nucleosides and bases are not directly synthesized de novo but are degradation products of nucleotides originating from for instance DNA and RNA.
The cell can reuse (salvage) nucleosides and bases, which can be transported
across the plasma membrane by facilitated diffusion or by concentrative
cotransport into the cell together with sodium (see “Nucleoside transporters”
below).
Nucleosides can be degraded to bases by purine nucleoside phosphorylase
(PNP), uridine phosphorylase or thymidine phosphorylase (TP). Purine bases
can be converted to NMPs by reactions catalyzed by adenine phosphoribosyltransferase (APRT) and hypoxanthine/guanine phosphoribosyltransferase
(HGPRT) (4).
In mammals, deoxynucleosides can be phosphorylated into dNMPs by the
deoxynucleoside kinases (dNKs) thymidine kinase 1 (TK1), thymidine
kinase 2 (TK2), deoxycytidine kinase (dCK) and deoxyguanosine kinase
(dGK) (5). Of these, TK2 and dGK are mitochondrial enzymes. The dNKs
are named from their best substrate but they also take other substrates. However they do not take ribonucleosides as substrates. Two mammalian ribonucleoside kinases (NKs) have been found in the cytosol, adenosine kinase
(AK) (6) and uridine/cytidine kinase (UCK) (7). No mammalian NK has
been found for guanosine and inosine phosphorylation.
Very importantly, nucleotides appear not to be transported across the
plasma membrane at significant rates. Therefore, the addition of the first
phosphate onto a nucleoside probably traps it inside the cell.
5’NTs catalyze the dephosphorylation of (d)NMPs. This reaction releases
the nucleoside to allow it to be transported out of the cell by facilitated diffusion. Seven human 5’NTs have been characterized so far (see “5’Nucleotidases” below).
Regulation of nucleotide metabolism
Types of regulatory mechanisms
The activity of some enzymes of the nucleotide metabolism is under tight
control. In general, enzymes can be regulated so that their catalytic rate,
localization or amount in the cell is changed. Often the first enzyme of a
pathway is regulated so that the cell can “save resources”.
However, as described above, metabolic pathways are often complicated
and branch at specific points into several pathways. Sometimes the ratio
between different end products has to be regulated, as with the dNTPs (described below). Many enzymes of nucleotide metabolism are regulated and
4
can together fine-tune the level of the various nucleotides in response to the
cell’s need.
All enzymes have an active site where they perform their catalysis. A basic form of regulation is by substrate availability. All enzymes need a minimum concentration of substrate in order to perform the catalysis, which is
dependent on how “tight” the substrate binds in the active site (the substrate
affinity).
Often enzymes are “feedback” inhibited by the end product of their pathway. This inhibition generally occurs when the end product reaches a concentration that exceeds the cell’s need. (d)NKs are feedback inhibited by
(d)NTPs (see below), which exert their inhibitory effect by binding in the
active site. Feedback inhibition can also occur through “allosteric” mechanisms.
Allosteric enzymes are regulated by specific “effector molecules” that
bind at a site that is separate from the active site. The binding of an effector
induces a conformational change in the enzyme that changes its catalytic
rate. This adds a way for nature to “turn on/off” enzymes. The cytosolic 5’nucleotidase II (cN-II) that is discussed in this thesis is an allosteric enzyme.
The mechanism for its allosteric activation is presented in Paper IV.
Some enzymes are also synthesized and degraded in a regulated manner,
as in the case of RNR (see below). There exist several more types of enzyme
regulation but that is beyond the scope of this thesis.
Regulation of de novo synthesis
The de novo synthesis of purine NMPs is largely regulated by allosteric
feedback inhibition by IMP, AMP and GMP. This regulates both the overall
rate of purine nucleotide synthesis but also the relative amounts of GMP and
AMP. The de novo synthesis of pyrimidine nucleotides is feedback inhibited
by CTP (1).
The de novo synthesis of dNTPs is mainly regulated by the allosteric enzymes RNR and dCMP deaminase. They control the ratios between the four
dNTPs (3, 4). RNR is allosterically regulated in a complex manner, in which
both its activity and substrate specificity is regulated through separate effector sites. It is activated by ATP and inhibited by dATP. Its specificity is
turned towards GDP, by the binding of dTTP; to ADP by the binding of
dGTP; and to CDP and UDP by the binding of dATP. Thus, it regulates the
ratios between dATP, dGTP and pyrimidine nucleotides (3, 8).
The ratio between dCTP and dTTP is regulated by dCMP deaminase that
converts dCMP to dUMP providing substrate for thymidylate synthase that
converts dUMP to dTMP (Fig. 4). dCMP deaminase is allosterically activated by dCTP and feedback inhibited by dTTP (3, 4).
5
Regulation of the salvage pathway
The salvage pathway is also regulated by feedback inhibition and allosteric
activation, which indicate its importance. dNKs and NKs are feedback inhibited by their corresponding dNTP, e.g. dCK is inhibited by dCTP and TK1 is
inhibited by dTTP (9-12).
The activities of the intracellular 5’NTs are probably largely dependent on
their substrate concentrations, since they have relatively low affinities for
their substrates. The 5’NTs cytosolic 5’-nucleotidase IA and IB (cN-IA and
cNI-B) are allosterically activated by preferably ADP, and cN-II is allosterically activated by preferably dATP and ATP. This reflects that they are sensitive to the energy state of the cell.
The 5’NTs and (d)NKs co-exist in the cell, catalyzing opposite and irreversible reactions. Thus they form “futile” substrate cycles together and can
thereby fine-tune the levels of phosphorylated nucleosides within the cell
(13) (Fig. 5). A single reversible enzyme would probably be less versatile in
this respect. In case of purine ribonucleotides, the 5’NTs indirectly oppose
the action of APRT or HGPRT.
Fig. 5. Substrate cycle between 5’NTs and (d)NKs. Nucleoside transporters (NTs)
mediate (deoxy)nucleoside transport across membranes.
Cell cycle dependence
The concentration of dNTPs in the cell varies during the cell cycle, being
highest during the DNA synthesis phase (S-phase). In resting cells, low
6
amounts of dNTPs are needed for DNA repair and mitochondrial DNA synthesis, which occur independently throughout the cell cycle. Many of the
anabolic enzymes of nucleotide metabolism are up-regulated before and
during the S-phase of the cell cycle and down-regulated in resting cells.
RNR of the de novo pathway consists of two subunits (R1 and R2). The
R2 is known to be degraded during mitosis and to be absent in resting cells
(14). Because of this it was also believed that the de novo pathway is primarily responsible for dNTP synthesis in proliferating cells and that the supply
of dNTPs in resting cells for cell cycle independent events such as DNA
repair and mitochondrial DNA synthesis is provided through the salvage
pathway. The salvage enzyme TK1 is also specific for the S-phase suggesting that the mitochondrial TK2 and dGK would be important in resting cells
(15).
Recently, a second form of R2 has been discovered that is expressed in
low amounts independently of the cell cycle. Since it can be induced by p53,
it was named p53R2 and proposed to provide the cell with dNTPs for DNA
repair (16). Mutation of p53R2 causes severe mitochondrial DNA depletion,
which indicates its importance for mitochondrial DNA synthesis (17). I was
shown that it is active in resting cells (18) and that resting cells have a complete de novo pathway (19). dTTP was formed in resting cells through the
action of an RNR consisting of R1 and p53R2 in the cytosol and by the salvage enzyme TK2 in mitochondria (20). The differential contributions to the
dNTP pools of dNTPs from the de novo pathway and the salvage pathway in
resting cells are not yet clear.
5’-Nucleotidases (5’NTs)
Overview
The first report of a 5’NT (EC 3.1.3.5) was made 1934 (21) and this family
of enzymes has been reviewed several times since then (22-24). They all
catalyze the hydrolysis of (d)NMPs into nucleoside and inorganic phosphate
(Pi) (Fig. 6). The human 5’NTs are listed in Table 1, assigned with aliases,
tissue specific expression and preferred substrates.
Fig. 6. 5’NTs dephosphorylates (d)NMPs.
7
Six of the seven characterized mammalian 5’NTs are intracellular enzymes that have similar fold and catalytic residues. These are the mitochondrial and cytosolic 5’(3’)-deoxynucleotidases (mdN and cdN), cN-IA, cNIB, cN-II and cytosolic 5’-nucleotidase III (cN-III). They work by a similar
catalytic mechanism, are completely dependent on Mg2+ for activity and are
inhibited by Pi.
The intracellular 5’NTs share the role of regulating intracellular nucleotide pools for maintaining balanced DNA and RNA synthesis, although they
also have more tissue-specific functions. Their Km values are very high relative to the substrate concentrations. However, their biological roles in for
instance regulation of dNTP pools are to some extent verified (see “Nucleotide pools” below).
The extracellular ecto 5’-nucleotidase (eN) has a different fold and catalytic mechanism relative the intracellular 5’NTs. It is anchored to the plasma
membrane via a glycosyl phosphatidylinositol (GPI) anchor. eN exhibits
high affinity for its substrates, especially AMP, and is transcriptionally regulated and participates in adenosine signaling.
8
Table 1. Properties of human 5’NTs
5’NT
Alias
Expression
Intracellular
mdN
dNT-2
cdN
dNT-1; PN-II;
UMPH-2
cN-IA
AMP-specific
5’NT; cN-I
cN-IA homologue; AIRP
High Km
5’NT; purine
5’NT; GMPIMP specific
NT
PN-I, P5’NT,
UMPH,
UMPH-1
cN-IB
cN-II
cN-III
Extracellular
eN
Ecto-5’NT,
NT5, eNT,
low 5’NT,
CD73
Preferred substrates, those
verified in vivo in bold
Ubiquitous
dUMP, dTMP, 3’dTMP,
3’UMP
Ubiquitous
dUMP, dTMP, dGMP,
dIMP,
3’dTMP, 3’UMP
Heart, skele- AMP, pyrimidine-dNMPs
tal muscle
AMP
Ubiquitous
GMP, IMP, dIMP, dGMP,
XMP
Phosphotransfer:
(deoxy)inosine
Erythrocytes CMP, UMP, dUMP,
dCMP, dTMP
Phosphotransfer:
Uridine, cytidine, deoxycytidine
Ubiquitous,
GPI anchored to
plasma
membrane
AMP, broad specificity,
both pyrimidine and purine
(d)NMPs
Cytosolic 5’(3’)-deoxynucleotidase (cdN)
cdN was first purified from human placenta (25) and later from erythrocytes
(26). It is ubiquitously expressed and shows highest catalytic efficiency
(Vmax/Km) for 3’dUMP, 3’dTMP, 3’UMP and 2’UMP, although the enzyme
also has high activity for all 5’dNMPs except dCMP (25, 26). Recombinant
murine cdN shows similar specificity as the human enzyme but also exhibits
some activity towards dCMP (27). cdN purified from erythrocytes was reported to have phosphotransferase activity (26), whereas cdN purified from
human placenta and the recombinant mouse cdN do not show phosphotransferase activity (25, 27).
9
Human cdN has a molecular mass of 23.4 kDa and is functional as a
dimer. The crystal structures of both human and murine cdN were solved by
us and are presented in Paper II.
cdN has mainly been studied for its role in regulating the cytosolic dTTP
pool, by dephosphorylation of dUMP and dTMP (see “Nucleotide pools”
below). It might also be important for catabolizing 3’- and 2’(d)NMPs produced by nucleases.
Mitochondrial 5’(3’)-deoxynucleotidase (mdN)
mdN shows 52% sequence identity to cdN and has similar size and oligomerization state as cdN. It is ubiquitously expressed and is targeted to the
mitochondria by a signal sequence. Recombinant human mdN prefers
dUMP, dTMP and 3’dTMP as substrates. However, it is not active with
purine nucleotides (28).
The crystal structure of human mdN was published together with a proposed catalytic mechanism for intracellular 5’NTs (29). Further structural
studies of this enzyme that involve its substrate specificity are presented in
Paper I and II.
Two inhibitors for mdN were identified (30):
(+)-1-trans-(2-Phosphonomethoxycyclopentyl)uracil (PMcP-U) a competitive inhibitor with a Ki of 40 μM, and
(s)—1-[2’-deoxy-3’,5’-O-(1-phosphono)benzylidene-b-D-threo-pentofuranosyl]-thymine (DPB-T), an inhibitor with mixed-type inhibition and a
Ki of 70 μM (30). PMcP-U is also a competitive inhibitor of cdN with a Ki
of 26 μM, while DPB-T is not. Crystal structures of mdN in complex with
PMcP-U and DPB-T showed the structural basis for the inhibition and proposed why DPB-T does not inhibit cdN (31). DPB-T can be used to distinguish between mdN and cdN and PMcP-U to distinguish between cytosolic
nucleotidases in enzyme assays (30).
The proposed function of mdN is to regulate the mitochondrial dTTP pool
by counteracting TK2 in a substrate cycle (see “Nucleotide pools” below).
Cytosolic 5’-nucleotidase IA (cN-IA)
cN-IA was first purified from pigeon heart where it shows high specific activity for AMP at millimolar concentrations (32). The enzyme is ~40 kDa
and appears to function as a tetramer (32). It is abundant in heart and skeletal
muscle (33, 34) but is also expressed in other tissues (35). Its structure has
not yet been solved.
Comparing catalytic efficiencies (Vmax/Km) the enzyme prefers deoxypyrimidine substrates over AMP (36). It is allosterically activated by
ADP and to a lesser extent by GTP (35, 37). A selective inhibitor for cN-IA
has been developed, 5-Ethynyl-dideoxyuridine (5-EddU), which is >100010
fold more selective for cN-IA than cN-II or eN, suggesting it is a useful tool
for specific inhibition of cN-IA (36, 38).
cN-IA might have a physiological function in heart in the generation of
signaling adenosine during ischemia (33, 39). It is activated by ADP and not
by ATP, which supports a role in AMP breakdown during ATP catabolism
(35-37). The high affinity towards dNMPs suggests a role in regulating the
dNTP pools.
Cytosolic 5’-nucleotidase IB (cN-IB)
cN-IB has four possible isoforms produced by alternative splicing, of which
the largest isoform has 610 amino acids. Counting 323 amino acids (identical between isoform 1-3), cN-IB has 67% sequence identity with cN-IA. It is
almost identical to the autoimmune infertility protein (AIRP) (40). It has the
highest expression in testis and its oligomerization state is not known. Its
structure has not yet been solved.
Overexpression of murine cN-IB in COS-7 cells showed that the enzyme
hydrolyzes AMP and is activated by ADP (40), but its substrate specificity is
still poorly characterized.
Cytosolic 5’-nucleotidase II (cN-II)
cN-II was first purified from chicken liver (41) and has been intensively
investigated in vertebrates for its specificity and regulation, but also for its
involvement in resistance towards anticancer drugs (see “Anti-viral/cancer
Nucleoside analogues” below).
It is ubiquitously expressed and functions as a tetramer (42-45). Recombinant human cN-II consists of 561 amino acids although the reported molecular mass of purified human cN-II varies. Its crystal structure was solved
by us and is presented in Paper III. A structural study of its substrate specificity and allosteric regulation is presented in Paper IV.
It prefers 6-hydroxypurine NMPs as substrates with the highest catalytic
efficiency for IMP and GMP followed by dIMP, dGMP and XMP (41, 4448). It possesses phosphotransferase activity (46-50), by which it can transfer a phosphate from a “donor” NMP (any substrate of the nucleotidase reaction) to an “acceptor” nucleoside, preferably inosine of deoxyinosine (51).
cN-II is allosterically activated by dATP, ATP, 2,3-bisphosphoglycerate
(2,3-BPG), polyphosphates and diadenosine polyphosphates (ApnAs) (43-45,
47, 52). The ApnAs are composed of two adenosines that are linked together
at the 5’-ends by a number of phosphates. Ap4-6As activate cN-II at micromolar concentrations. These are formed as byproducts of aminoacyl tRNA
synthetases and are removed by ApnA hydrolases before they reach toxic
levels (53, 54). They have been reported as both intra- and extracellular signaling molecules (55).
11
The enzyme might have a complex regulation involving a second effector
site (56, 57) (Paper III) and its 13 residue C-terminal acidic stretch that
might mediate regulation via subunit association/dissociation (45).
cN-II appears to be involved in regulating the IMP and GMP pools within
the cell (see “Nucleotide pools” below), and indirectly also participate in
regulating the AMP pool.
Increased cN-II levels were found in patients suffering from Lesch-Nyhan
syndrome, which is caused by a deficient HGPRT (see “Nucleotide pools
and disease” below). Possibly cells with deficient HGPRT then instead use
the phosphotransferase activity of cN-II to produce GMP.
Recently, knockdown of cN-II by RNA interference was shown to lead to
apoptosis in astrocytoma brain tumor cells (58).
Cytosolic 5’-nucleotidase III (cN-III)
cN-III has mainly been studied in red blood cells but has also been found in
other tissues (59-61). It functions as a monomer (26, 61). The cN-III gene
gives rise to four alternatively spliced mRNA transcripts, predicted to produce enzymes of the lengths 297 (isoform 1), 286 (isoform 2), 285 (isoform
3) and 336 (isoform 4) amino acids. Isoform 2 (Q9H0P0-2) is identical to the
p36 protein that has been found in inclusion bodies in lupus and AIDS (60).
cN-III shows highest catalytic efficiency (Vmax/Km) towards CMP followed by UMP, dCMP, dUMP and dTMP, and does not take purine nucleotides as substrates (62). Similar to cN-II, this enzyme has phosphotransferase
activity. The transferred phosphate can originate from any monophospate
substrate, whereas uridine and cytidine are shown to be the best phosphate
acceptors (26).
cN-III is believed to catabolize UMP and CMP produced by RNA degradation, in red blood cells during the final steps of erythroid differentiation
(63). During these steps, the RNA content of red blood cells is drastically
reduced and cN-III appears to be up-regulated (63, 64).
A reduction of cN-III activity due to genetic defects or lead poisoning
causes hemolytic anemia in humans (65). Here the circulating red blood cells
show significant accumulation of pyrimidine nucleotides as well as incomplete degradation of RNA and ribosomes, which indicate the important role
of cN-III in red blood cell (63).
19 different mutations related to hemolytic anemia have been identified so
far (66, 67). Many of these, e.g. the missense mutations giving D87V,
L131P, N179S, G230R, C63R, G157R and I247T reduce the activity or stability of the enzyme (61, 64, 67-70) (isoform 2: Q9H0P0-2).
The crystal structure of murine cN-III has been published (71) and the
structure of human cN-III was solved by us and is discussed in this thesis.
These structures display where the substitutions are located, which enlights
how they could affect protein function.
12
Ecto-5’-nucleotidase (eN)
eN has a different fold and catalysis relative the intracellular 5’NTs, with a
catalysis depending on a dimetal center (see “Catalytic mechanism of eN”
below). It is attached extracellularly to the plasma membrane by a GPI anchor. The mature enzyme is of 548 residues and has a calculated molecular
mass of 61 kDa and functions as a dimer.
The crystal structure of an E. coli homologue of human eN has been
solved (72). A homology model was made of the human eN based on the E.
coli structure (73).
eN has a broad substrate specificity, although AMP is the most efficient
substrate considering Vmax/Km values. The Km values for AMP are in the low
micromolar range and ADP and ATP are competitive inhibitors. Because of
the high affinity for AMP and the fact that extracellular adenine nucleotides
are relatively abundant, AMP has been considered as the major substrate.
Extracellular adenosine, ADP, ATP and some other nucleotides function
as signaling molecules for purinergic receptors. Adenosine acts on adenosine
(P1) receptors and eN probably has a biological role in releasing adenosine
for adenosine receptor activation (74-78).
eN is expressed in most tissues although only in certain cell types within
each tissue (79). This is controlled at the transcriptional level and several
transcription factor-binding sites are present in the promoter region that may
mediate both positive and negative regulation. Ischemia and hypoxia has
been shown to induce eN expression (80-82), indicating a role for eN in signaling events related to these conditions.
eN probably also affects the intracellular nucleotide pools, since it’s nucleoside products can pass the plasma membrane through nucleoside transporters.
Nucleotide pools and disease
The size of the dNTP pools and their relative amounts are important. It was
early demonstrated in E. coli that depletion of the dTTP pool induces cell
death (83). The consequences of imbalanced dNTP pools are also reflected
in several genetic diseases that are associated with deficiencies in enzymes
of nucleotide metabolism.
Deficiencies in adenosine deaminase (ADA; that converts deoxyadenosine and adenosine to deoxyinosine and inosine) or in PNP lead to
sever immune deficiency due to defects in T and B cell function. These defects are associated with accumulation of dATP and dGMP, respectively (84,
85). Imbalance in dATP or dGTP pools affect the activity of RNR, thus affecting also the other dNTP pools. Elevated levels of dATP might trigger
13
apoptosis, since dATP participates together with cytochrome c in triggering
apoptosis through Apaf-1 mediated activation of procaspase 9 (86, 87).
Deficiency in the salvage enzyme HGPRT, which causes Lesch-Nyhan
syndrome, leads to accumulation of hypoxanthine and guanine followed by
overproduction of uric acid, which causes problems such as severe gout,
poor muscle control, and moderate mental retardation (88).
Deficiency of TP that degrades thymidine and deoxyuridine has been
found in patients with the mitochondrial neurogastrointestinal encephalomyopathy (MNGIE), which is a disease linked to e.g. deletions in mitochondrial DNA (89). This causes mitochondrial dysfunction that gives severe
symptoms (90). Increased levels of thymidine and deoxyuridine (from <0.5
μM to 10-20 μM) were found in blood plasma of patients with MNGIE (91).
This leads to elevation of cytosolic and mitochondrial dTTP and to other
effects that cause imbalance in dNTP pools.
Loss-of-function mutations in TK2, dGK and in the RNR subunit p53R2
(see below) have been found in patients suffering from mitochondrial DNA
depletion syndrome (MDS) (17, 92-94). MDS causes quantitative mitochondrial DNA depletion. The concomitant low amount of mitochondrial DNA is
presumably causing insufficient synthesis of the mitochondrial DNAencoded proteins involved in oxidative phosphorylation, causing loss of
muscle activity and death in early childhood (90, 92).
The only 5’NT deficiency known is that of cN-III, which has been associated with hemolytic anemia (discussed above) (65).
Nucleotide pools
The dNTP pool sizes within the cell are rather small and vary during the cell
cycle (Table 2). During the S-phase of the cell cycle these pools would be
diminished within a few minutes without biosynthesis. The genetic diseases
discussed above underline the importance of maintaining balanced nucleotide pools. The dNTP pools are much smaller in resting cells than in differentiating cells (95). The cytosolic and mitochondrial pools of dTTP and
dGTP are exchanged rapidly and share a dynamic equilibrium (96).
14
Table 2. Average dNTP (97) and NTP (98) pool sizes in mammal cells
(d)NTP
dTTP
dATP
dCTP
dGTP
UTP
ATP
CTP
GTP
Mitochondria
Cytosol
6
pmole/106 cells (μM if
pmole/10 cells (μM if
106 cells is 0.02 μl)
106 cells is 0.2 μl)
0.88 (44)
32 (160)
0.75 (37.5)
17.5 (87.5)
0.74 (37)
27.5 (137,5)
0.40 (20)
5.9 (29.5)
Concentrations below are shown in μM
480
2800
210
480
In situ studies on the substrate cycles between dNKs and 5’NTs have provided evidence supporting that dNKs and 5’NTs are involved in the regulation of specific nucleotide pools. Mammal cell lines lacking a dNK or overexpressing a 5’NT were produced (99, 100). By measuring excretion of nucleosides from these cell lines by isotope flow experiments, the involvement
of dNKs and 5’NKs could be measured (97, 100-102).
This provided evidence for anabolic roles of TK1 in thymidine/TMP and
deoxyuridine/dUMP substrate cycles and of dCK in deoxycytidine/dCMP
and deoxyadenosine/dAMP substrate cycles (99, 103).
cN-II was indicated not to be involved in the regulation of the deoxynucleotide pools but instead affect the IMP and GMP pools (100). Recombinant murine cdN gave evidence for a catabolic role of cdN in substrate cycles regulating pyrimidine nucleotide pools (100). Early studies on human
cN-III deficient erythrocytes showed that dTMP and dUMP, but not UMP,
CMP and dCMP were degraded in these cells. This indicated the existence of
another pyrimidine specific cytosolic 5’NT, besides cN-III, that took dTMP
and dUMP, but not UMP, CMP or dCMP as substrates (59, 104). This enzyme was later identified as cdN.
It was also demonstrated that mdN participates in a substrate cycle together with TK2 in regulating the mitochondrial dTMP pool (97).
There are still many gaps in the understanding of the cellular functions of
the different 5’NTs. For instance, it is yet not clear which 5’NTs regulate the
cytosolic or mitochondrial dGTP or dATP pools. Although for regulation in
the cytosol, in vitro kinetic studies suggest cdN and cN-II for dGTP regultation and cN-IA for dAMP regulation.
15
Nucleoside/nucleotide transporters
Two types of mammalian nucleoside transporters have been characterized.
These are the equilibrative nucleoside transporters (ENTs) that transport
nucleosides and bases through the plasma membrane bidirectionally down
their concentration gradient, and the concentrative nucleoside transporters
(CNTs) that cotransport nucleosides against their concentration gradient into
the cell together with sodium. The CNTs use the force derived from transporting sodium down its concentration gradient to concentrate nucleosides
inside the cell (105).
Of the four known human ENTs (hENT1 (106), hENT2 (107), hENT3
(108) and hENT4 (109), hENT1 and hENT2 are mainly found in the plasma
membrane and have broad tissue distribution. They transport both
pyrimidine and purine nucleosides and ENT2 can also transport some bases.
These two can be distinguished by their sensitivity to inhibition by nanomolar concentrations of nitrobenzylmercaptopurine ribonucleoside (NBMPR),
hENT1 being sensitive and hENT2 being insensitive (105). ENT1 has also
been found in membranes of mitochondria, lysosomes, the nuclear envelope
and the endoplasmic reticulum (105).
Three human CNTs (hCNT1, hCNT2 and hCNT3) have been characterized (110-112). The CNTs are highly expressed in intestine, kidney and
liver. This implies that they are involved in absorption and excretion of dietary nucleosides (105). ENTs and CNTs are important for the cellular uptake
of drugs used in the treatment of cancer and viral diseases (105).
When nucleosides become phosphorylated into nucleotides, they can no
longer be transported across the plasma membrane. However, they can diffuse through nuclear pores and there is no concentration gradient of nucleotides between the cytosol and the nucleus (113). Nucleotides are transported
in a controlled manner in and out of the mitochondria. The adenine nucleotide carrier (ANC) catalyzes transport of ADP into mitochondria in exchange
for ATP, which is needed for the generation and distribution of ATP, that is
formed from ADP by the action of ATP synthase within the mitochondria.
A mitochondrial carrier (SLC25A19) was reported as a deoxynucleotide
carrier (DNC) that catalyzes transport of (d)NDPs and (d)NTPs across the
mitochondrial inner membrane (114). Later studies indicate that SLC25A19
is not a deoxynucleotide carrier, but might be a carrier for thiamine pyrophosphate (115-117).
A highly specific import system for dTMP into the mitochondria was
demonstrated (118). This could be the main road for dTTP supply to mitochondria, since dTMP is the first thymidine nucleotide formed during de
novo synthesis (118). Evidence for transport of dCTP and UTP in mitochondria have also been reported (119, 120).
16
Anti-cancer/viral nucleoside analogues
Nucleoside analogues (NAs) are widely used in treatment of cancer and viral
diseases. Fig. 7 provides chemical structures and names of some NAs. Some
of these have been used in treatment of HIV-1 (e.g. AZT and d4T), or other
viral diseases (e.g. BVDU and ribavirin). Others are currently used in treatment of cancer (e.g. araC, dFdC and CdA). NAs are prodrugs that need to be
phosphorylated within the cell to become active. They are given to the patient as nucleosides since their phosphorylated forms cannot pass through the
plasma membrane.
To exert their cytotoxic effect, many NAs are phosphorylated to their
triphosphate form by (d)NKs, NMPKs and NDPKs (5, 121, 122) and then
presumably incorporated into DNA leading to DNA chain termination.
Many NAs also act through enzyme inhibition, e.g. inhibiting DNA polymerase α or RNR (123). Inhibition of RNR decreases the biological dNTP
pools and thereby increases the NA-TP/dNTP ratio.
17
Fig. 7. Nucleoside analogues. Top: AZT: 3’-azidothymidine, BVdU: E)-5-(2bromovinyl)-2’-deoxyuridine and d4T: 2’,3’-didehydro-2’,3’-dideoxythymidine.
Middle: AraC: 1-β-D –arabinofuranosylcytosine, dFdC: 2’,2’-difluoro-2’deoxycytidine (Gemcitabine) and CdA: 2’-chlorodeoxyadenosine. Bottom: Ribavirin.
A successful NA treatment is highly dependent on the level of cell uptake
through plasma membrane transporters and on how efficiently they are
phosphorylated into their active form (105, 123).
5’NTs can dephosphorylate the monophosphate form of many NAs (NAMPs). This is not only a step in the inactivation of the drugs, but it also releases them to exit the cell.
Patients often evolve resistance to NAs. The mechanism behind this has
been studied intensively, e.g. concerning resistance to cytosine and adenine
18
based NAs such as araC, dFdC and CdA in patients suffering from acute
myeloid leukemia (AML). 5’NTs, ENT1, RNR, dCK and other enzymes
involved in metabolism or uptake of NAs have been the focus in these studies (123).
Several studies link increased cN-II activity or mRNA expression with resistance to cytosine and adenine based NAs in cell lines and poor clinical
outcome of araC treatment of AML patients (124-126). Inhibition of cN-II
resulted in a 2-5 -fold increase in dFdC cytotoxity in lymphocytic Bleukemia cells (127). However, cN-II expression was not linked to dFdC
resistance in patients with advanced non-small cell lung cancer: here high
levels of cN-II mRNA instead correlated with increased survival (128).
The focus on cN-II in studies of adenine and cytidine based NAs can
seem somewhat strange since it has been known for a long time that cN-II
has very low activity towards cytosine and adenine based nucleotides. More
recent in vitro activity measurements on recombinant cN-II indicated a very
low activity towards such NA-MPs (Table 5) (30). This was supported by
the observation that increased expression of cN-II in human 293 cells gave
no increased resistance towards toxicity of CdA (129). However, dephosphorylation of the monophosphophate form of fludarabine (an adenine-based
NA; F-ara-AMP) by cN-II was measured by HPLC and NMR (130), which
suggested a role of cN-II in dephosphorylating this analogue.
A 5’NT with known high activity towards AMP is cN-IA. Overexpression
of cN-IA in human cell lines generated resistance towards CdA and dFdC
(35, 131). cN-IA dephosphorylated CdAMP with a specific activity of 39%
compared to that of AMP (35).
cN-II might activate guanosine and inosine analogues by its phosphotransferase activity. cN-II was suggested as the main enzyme activating
the anti-HIV analogue 2’,3’-dideoxyinosine (ddI) (50) . It was also shown to
activate the antiviral NA ribarivin significantly better than the human
(d)NKs (132).
cdN could in principle be involved in resistance towards thymine, uracil
or guanine based NAs, considering its substrate specificity and ubiquitous
expression. It also shows some activity towards such NAs in vitro (Table 5)
(30).
The mitochondrial mdN is however not likely to be involved in NA resistance, since the NAs are mainly targeted to nuclear DNA. Instead mdN
might affect the mitochondrial toxicity that is commonly experienced as a
side effect of NA treatment. The mechanism behind this toxicity comes from
competitive inhibition of mitochondrial enzymes such as the mtDNA polymerase γ. Such inhibition of the mtDNA polymerase γ, by AZT, was associated with mtDNA depletion in HIV-1-infected patients (133).
19
Evolution and catalysis of intracellular 5’NTs
Overall structure of intracellular 5’NTs
The intracellular 5’NTs belong to the haloacid dehalogenase (HAD) superfamily, which includes phosphohydrolases such as the 5’NTs, phosphotransferases, P-type ATPases, phosphonatases, dehalogenases and sugar phosphomutases (134, 135).
The HAD superfamily is characterized by a core domain, consisting of an
α/β-Rossmannoid fold, containing a three-layered α/β sandwich built up
from repeating β−α units. The β−sheet is parallel and consists most often of
at least five strands in a 54123 strand order, which is typical for the α/βRossmann fold.
The fold of HAD superfamily enzymes can be distinguished from other
α/β-Rossmann-like folds by two structural signatures, a “squiggle” (a single
helical turn) and a “flap” (a β-hairpin motif) (135). The “squiggle” is located
immediately downstream of strand S1 and the “flap” is located downstream
of the “squiggle” (Fig. 8a).
Most HAD superfamily enzymes also contain a cap domain, which is
much less conserved than the core domain. This cap domain is inserted either (1) in the middle of the β-hairpin of the “flap” motif or (2) immediately
after strand S3. mdN has the cap domain inserted after the “flap” motif (Fig.
8a). Bioinformatic analysis (135) indicates that the cap domain has been
inserted at multiple occasions during evolution, independently of the core
domain, and that this has enabled the great diversity in function of these
enzymes.
The active site is located in the cleft between the core and the cap domains with the catalytic residues in the core domain and most of the residues
that determine substrate specificity in the cap domain. In the 5’NTs, the substrate binds with its phosphate and ribose moieties in the core domain and its
base moiety in the cap domain (Fig. 8b). The cap domain probably determines the base specificity of the 5’NTs.
Most of HAD enzymes share four structurally conserved motifs that take
part in forming the catalytic Mg2+-phosphate site (with some exceptions, e.g.
dehalogenases). Motif I (DXDX[T/V]) is located on strand S1 and on the
downstream “squiggle” loop, and motif II ([T/S]) on strand S2. The locations of motif III ([K/R]) and motif IV (DX0-4D) are less conserved. Motif III
is found on the loop downstream of strand S3 and motif IV is located in
strand S4 and downstream of it. (motif III and IV are sometimes described as
both included in motif III) (Fig. 8b).
20
Fig. 8. HAD superfamily fold and conserved motifs (D41N variant of mdN; Paper
I). a) The core domain (light) and cap domain (dark), with strands S1-S5, squiggle
(Sq) helical turn, Flap β−hairpin, C-terminal (C) and N-terminal (N). b) Motif I
residues: N(D)41, D43 and V45. Motif II residue: T130. Motif III residue: K165.
Motif IV residues: D175, D176. dUMP and Mg2+ are also shown.
The annotation of intracellular 5’NTs to the HAD superfamily was first
proposed by sequence analysis (136), which was later confirmed by the
structure of human mdN (29), followed by those of human cdN (Paper II),
human cN-II (Paper III) and human cN-III (unpublished). The murine cdN
was also solved by us (Paper II), whereas the murine cN-III structure was
published by another group (71).
cN-II has a large cap domain following the “flap” motif and an additional
fold following strand S3 (Fig. 9).
cN-III is structurally very similar to phosphoserine phosphatase (PSP),
which dephosphorylates phosphorylated L-serine (137). Both cN-III and PSP
have cap domains following the “flap” motif, but also small contributions to
the cap domain inserted after strand S3 (Fig. 10).
21
Fig. 9. cN-II structure with cap domain (dark) and core domain (light). Squiggle
(Sq) motif and Flap motif are marked.
Fig. 10. Overall structure of a) cN-III and b) PSP, with cap domains (dark) and core
domain (light). Squiggle (Sq) motif and Flap motif are marked, as well as strands
S1-S5.
22
Catalytic mechanism
The catalytic mechanism of intracellular 5’NTs (Fig. 11) was proposed
based on the structures of mdN that mimic reaction intermediates 3, 4 and 6
of the proposed mechanism (29). It has been supported by a theoretical study
(138) and is also very similar to the catalytic mechanism proposed for PSP
(139). The catalytic mechanism of 5’NTs starts as follows:
The first aspartate of motif I initiates the catalytic reaction by making a
nucleophilic attack on the phosphate moiety of the substrate. Hypothetically,
it might form a pentavalent substrate intermediate (Intermediate 2) (140).
This nucleophilic attack is probably conserved throughout the HAD superfamily.
The second aspartate of motif I appears to be bifunctional. It first acts as a
general acid, donating a proton to the leaving nucleoside product. A phosphoenzyme intermediate probably remains (Intermediate 3 and 4) (136, 141).
This Asp then probably acts as a general base, activating the water molecule
that then hydrolyses the phosphate (Intermediate 5 and 6).
The second Asp of motif I is present in most HAD superfamily enzymes
where there is a need for protonation of the leaving group. It is not present in
for instance HAD (in which the first leaving group is a Cl- ion) or in P-type
ATPases (where the leaving group is ADP) (Table 3). Neither Cl- nor ADP
need to take up a proton. The threonine that substitutes the second Asp in Ptype ATPases appears to have a function in stabilizing the phosphoenzyme
intermediate, enabling the necessary conformational change that drives the
ion transport.
Motif II (Thr/Ser) is very conserved and appears to coordinate the phosphate moiety in many HAD enzymes. Motif III (K/R) is also very conserved.
It probably has a role in stabilizing negative charge in reaction intermediates.
Motif IV contains two aspartates, of which one is coordinating the Mg2+
and the other has an unclear role. The sequential order of these two aspartates varies in different enzymes. In Table 3, the Asp with unknown role is
marked Asp*.
We speculated in Paper IV over the role of this Asp*. It is clearly involved in the mechanism of allosteric activation of cN-II. This Asp* is very
conserved among the HAD enzymes. Substitutions of this residue abolish the
enzyme activity in mdN, PSP, HAD and Sarcoplasmic Ca2+-ATPase
(SERCA) (142-144 and unpublished data on mdN).
23
Fig. 11. Proposed catalytic mechanism of intracellular 5’NTs. The scheme is taken
from reference (29).
24
Table 3. Conseved motifs of HAD superfamily enzymes
Protein (pdb code) Motif I
Motif II Motif III
DXDX[V/T]
[T/V]
[K/R]
C–Cl cleavage
HAD (1QQ6)
8DAYGT
S114
K147
C–P cleavage
PALD (1FEZ)1
12DWAGT
T126
R160
CO–P cleavage
PSP (1F5S)
11DFDST
S99
K144
Human mdN
41DMDGV
T130
K165
Human cdN
10DMDGV
S99
K134
Human cN-IA
211DGDAV Human cN-IB2
467DGDAV Human cN-II
52DMDYT
T249
K292
Human cN-III3
38DFDMT
S153
K202
MPP (1XVI)4
13DLDGT
S47
K188
MDP1 (1U7O)5
11DLDYT
S69
K100
CO–P cleavage/formation (mutase)
8DLDGV
S114
K145
β-PGM (1O03)6
CO–P cleavage/formation (mutase)
SERCA (2ZBD)
351DKTGT
T625
K684
H-pump (3B8C)7
329DKTGT
T511
K569
8
Na/K ATPase
369DKTGT
T610
K691
1
Phosphonoacetaldehyde hydrolase
2
Isoform 1 (Q96P26-1)
3
Isoform 2 (Q9H0P0-2)
4
Mannosyl 3-phosphoglycerate phosphatase
5
Mg2+-Dependent phosphatase 1
6
β-phosphoglucomutase
7
Plasma membrane proton pump from Arabidopsis thaliana
8
Na/K ATPase from pig (3B8E)
Motif IV
DX0-4D
D214, D218*
D186, D190*
D167, D171*
D175*, D176
D144*, D145
D351, D356*
D227, D231*
D214, D218*
D122*, D123
E169*, D170
D703, D707*
D588, D592*
D710, D714*
Catalytic mechanism of eN
The extracellular eN dephosphorylates (d)NMPs by a mechanism different
from that of the intracellular 5’NTs. The structure of the E. coli homologue
of eN showed that it belongs to a large superfamily of metallophosphoesterases that use a dimetal center for catalysis (72).
Based on the structure of E. coli eN in complex with α,β-methyleneADP, a catalytic mechanism was proposed (145). The E. coli eN has broader
substrate specificity than human eN, but consists of very similar structural
motifs participating in the catalysis. The E. coli eN can dephosphorlyate not
25
only monophosphates but also di- and triphosphates, whereas the human
enzyme is competitively inhibited by di- and triphosphate nucleosides.
One oxygen atom of the terminal phosphate group of α,β-methylene-ADP
coordinates the dimetal centre. A catalytic histidine and an arginine were
proposed to polarize the phosphate group to prepare it for the nucleophilic
attack, which appears to be carried out by a water molecule (145).
The mechanism of E. coli eN also involves a large domain rotation (146,
147). This might be conserved to human eN, although a homology model of
human eN indicates that the active site is more accessible due to a shorter
loop (73).
26
Present Investigation
Aim
The focus of my PhD project was to study substrate recognition, catalysis
and regulation of 5’NTs. The main method used was x-ray crystallography.
The project was initiated with the aim to solve structures of mdN and cdN
in complex with substrates. We wanted to analyze their substrate recognition
in detail, which would increase the understanding of their substrate specificity and their possible involvement in the degradation of NA-MPs.
The project continued with the aim to determine structures of the remaining human intracellular 5’NTs. We solved the structures of human cN-II and
human cN-III. We aimed at investigating the structural basis for their substrate specificity and to study the allosteric regulation of cN-II.
Strategy
To achieve our aims, we needed to produce crystals of the enzymes with
substrate (or nucleoside product) bound in the active sites.
This might seem straightforward at first glance, but substrate/product
complexes cannot be obtained unless the enzyme can be stabilized in a state
where the substrate/product can stay bound for a longer time.
A reason for this is that enzymatic reactions such as the hydrolysis of a
phosphomonoester bond often are extreamly fast and substrate/product intermediates very short-lived (148). Crystal structures are built from data that
reflect the “average” state of the protein; the structures show the reaction
intermediate that dominates at the equilibrium of the crystal. Because of this,
it is very unlikely to obtain a structure of a 5’NT in complex with substrate
or nucleoside product, without first performing a trick. We used two strategies to produce substrate/nucleoside product complexes, see below. Several
structures of various complexes were solved (Table 4).
27
Fluorometallic complexes as models for enzyme
intermediates
Fluoride has long been known to be very toxic. In complex with metals,
fluoride is known to inhibit nucleotide-binding proteins, e.g. phosphatases,
phosphorylases and many ATPases, and also activate G-proteins participating in signal transduction. It has a preference to form complexes with metals
such as aluminium, beryllium or magnesium. These complexes, e.g. BeF3-,
MgF3- and AlF4-, mimic phosphate and bind with high affinity to phosphate
sites in proteins. They also form tight penta- or hexavalent complexes with
reaction products, trapping them in the active site thus locking the enzyme in
a state similar to e.g. a substrate intermediate state (149).
This property of fluorometallic complexes has been used extensively by
crystallographers to mimic reaction intermediates, in order to study catalytic
mechanisms. Such studies were made on HAD superfamily enzymes e.g.
mdN (29) and PSP (139) and several other proteins e.g. cAMP-dependent
protein kinase (150) and transducin α (151). This strategy was used several
times in the present investigation, e.g. we solved the structure of human cdN
in complex with the product deoxyuridine together with AlF4- (Paper II).
It is highly unlikely to capture the “real” high-energy reaction intermediate of a phosho-transfer reaction. In spite of this, a structure of β−PGM in
complex with a “real” pentacoordinated phosphorane intermediate was published (140). This intermediate was later indicated by 19F NMR to be a
MgF3- complex (152). Recently, it was also shown by 19F NMR that tetrahedral fluorometallic complexes modeled in electron densities as AlF3- are
probably MgF3- (153).
Substitutions to trap substrate in active site
The proposed catalytic mechanism of 5’NTs helped us to design inactive
variants of mdN (D41N), human cdN (D10N), murine cdN (D12N), cN-II
(D52N), and cN-III (D38N, isoform 2: Q9H0P0-2), with the purpose of
binding substrates in the active site without hydrolysis. In these variants, the
first aspartate of motif I (which in the catalytic reaction makes a nucleophilic
attack on the phosphate of the substrate) was mutated into an asparagine.
This asparagine forms a strong hydrogen bond to the phosphate probably
further increasing the affinity for the substrates without hydrolysis.
Several structures of human mdN (D41N) and murine cdN (D12N) in
complex with various substrates and NAs were determined (Paper I and II).
The cN-II variant D52N enabled us to bind substrates together with activators (Paper IV).
We could not obtain structures of substrate complexes of the D10N variant of human cdN, probably since Pi (present in the crystallization solution)
28
competively inhibited the substrate to bind. The crystals did not survive
longer soaking experiments for the necessary removal of Pi.
Table 4. Structures solved
Protein
Wild type mdN
D41N variant of mdN
Wild type human cdN
D10N variant of human cdN
Wild type murine cdN
D12N variant of murine cdN
Wild type cN-II
D52N variant of cN-II
Wild type cN-III
1
Structures (Resolution in Å;pdb code)
FdU-AlF4- (1.7)1
Ca2+ (1.9)1
dUMP (2.0; 1Z4I)2
dTMP (1.8; 1Z4L)2
dGMP (2.0; 1Z4P)2
UMP (1.7; 1Z4M)2
3’dTMP (1.75; 1Z4K)2
2’UMP (1.8; 1Z4J)2
d4TMP (2.05; 1Z4Q)2
AZTMP (1.8; 2JAU)3
BVdUMP (1.95; 2JAW)3
PO4 (1.4)1
Deoxyuridine-AlF4- (1.2; 1SRW)3
PO4 (1.05)1
Deoxyuridine-BeF3- (1.5)1
Thymidine-AlF4- (1.6)1
dUMP (1.9; 2JAR)3
dGMP (2.0; 2JAO)3
Ado (1.5; 2JC9)4
BeF3 (2.15; 2JCM)4
SO4 (2.2; 2J2C)4
Ap4A-SO4 (2.2)1
Apo (2.3)5
ATP (2.0)5
IMP-ATP (1.9)5
IMP-BPG (2.3)5
GMP-Ap4A (2.0)5
dGMP-dATP (2.3)5
UMP-ATP (2.3)5
GMP-BPG (2.5)1
Carboxy-Asp-BPG (2.1)1
Apo (2.7; A2CN1)1
BeF3 (2.5; 2VKQ)1
PO4 (3.0; 2JGA)1
Unpublished, 2Paper I, 3Paper II, 4Paper III, 5Paper IV
29
Structural basis for substrate specificity of mdN
and cdN (Paper I and II)
mdN and cdN share 52% sequence identity. They differ in their substrate
specificity (Table 5), mdN being very specific for the uracil and thymine
based nucleotides, whereas cdN also takes dGMP and dIMP as substrates.
Murine cdN is 85% sequence identical to human cdN. Human and murine
cdN have similar specificities although the murine enzyme has some activity
also for dCMP (Table 5). We wanted to characterize the structural basis for
the substrate specificity of mdN and cdN, and to analyze the basis for the
differences between them in substrate specificity.
The three enzymes have very similar overall structures and their active
sites are also very similar. Despite this, they have some important differences in the base recognition site of the cap domain: Residues Ile133, Trp76
and Trp96 in mdN correspond to Leu45(47), Tyr45(47) and Leu102(104) in
human and murine cdN (the residue numbers in parenthesis refer to murine
cdN).
30
Table 5. Substrate Specificities of cdN, mdN and cN-II
Relative enzyme activity, %
Substrate
Human cdNa Murine cdNb Human mdNc Human cN-IId
dUMP
100
100
100 (100)
23
dTMP
55
65
50 (48)
16
dCMP
0.7
16
0 (0)
1
dAMP
13
9
2 (1)
7.5
dGMP
81
45
6 (2)
72
dIMP
245
96
8 (3)
86
21
11
Nd
IMP
100
GMP
4.5
4
2 (0)
85
AMP
0
1
0 (1)
7
Nd
Nd
Nd
XMP
23
UMP
8.9
16
30 (37)
14
CMP
0
0
0 (0)
6
Nd
58
77 (103)
3’dTMP
Nd
3’UMP
76*
35
47 (34)
Nd
2’UMP
53*
11
18 (15)
Nd
Nucleoside
analogue
4
3
9
d4TMP1
0.6
22
103
2
AZTMP1
6
8.5
19
28
BvdUMP1
1.5
217
108
82
FdUMP1
11
0.1
Nd
1
AraTMP1
0.2
1
18
0.1
ddCMP1
1
2
20
0.1
dFdCMP1
0.6
0.1
0.2
0.1
araCMP1
0.2
1.5
1.2
0.2
3TCMP1
0.2
0.5
Nd
0.1
CdAMP1
4.5
1
Nd
0.3
araGMP1
3.5
The relative enzyme activities are taken from previous publications:
1
(30)
a
(25) (*calculated from the Vmax values)
b
(27) (in parenthesis is measured at pH 7.5) c(28) which also report Km and
Vmax values for the individual enzymes.
d
(44)
Nd: Values not found in literature
31
Pyrimidine base specificity
The base recognition site of both mdN and cdN contains two main chain
amides that form favorable hydrogen bonds to the 4-carbonyl group of
(d)UMP and (d)TMP, but would repel the amino group of (d)CMP. In cdN,
the two main chain amides also favorably bind the 6-carbonyl group of
(d)GMP and (d)IMP, but would repel the amino group of (d)AMP (Fig. 1,
Fig. 12).
In this way, mdN and cdN can discard cytosine and adenine based nucleotides as substrates (Paper I and II).
Fig. 12. mdN-dUMP and 4-carbonyl specific hydrogen bonds to main chain amides.
32
Deoxy/ribo specificity
Both mdN and cdN prefer the deoxy-form over the ribo-form of nucleotides
(Table 5). They accomplish this specificity by having a hydrophobic patch
stacking the 2’-C of deoxy-substrates (Fig. 1). Fig. 13 shows the mdNdUMP structure and the mdN-UMP structure, superimposed.
The 2’C of dUMP is favorably stacked by Phe49, Phe102 and Ile133,
while the 2’OH of UMP binds unfavorably close to these residues, with the
closest distance (2.7 A) to Ile133. Clearly, the hydrophobic patch of Phe49,
Phe102 and Ile133 provides the deoxy specificity of mdN (Paper I). This
hydrophobic patch is also conserved in cdN, where the corresponding residues in human cdN are Phe18, Phe71 and Ile102.
Fig. 13. mdN-dUMP structure (light) and mdN-UMP structure (dark) superimposed.
Dotted lines indicate the unfavorable interactions between the 2’OH and hydrophobic/aromatic residues.
33
2’-,3’- and 5’-phosphate specificity
The relatively high activity of mdN and cdN towards 2’- and 3’phosphorylated (d)NMPs made us compare the substrate recognition of them
with the 5’dNMPs (Fig. 14) (Paper I).
The phosphate moieties are recognized similarly, whereas the ribose and
base moieties of 3’dTMP and 2’UMP are rotated relative in dUMP (Fig. 14).
The bases of 3’dTMP and 2’UMP appear to be more favorably stacked by
the aromatic Phe49, than the pyrimidine base of 5’substrates (Fig. 14). This
supports the idea that at least 3’-phosphorylated pyrimidine (d)NMPs are
biologically relevant substrates.
It becomes clear that 3’- and 2’-purine NMPs would probably not bind
with high affinity, since the purine base probably would clash into aromatic
residues of the active site.
Fig. 14. mdN in complex with dUMP, 3’dTMP and 2’UMP superimposed.
34
Purine/pyrimidine specificity
Human and murine cdN have a wider base recognition site than mdN, which
probably is the basis for why they have higher activity for purine nucleotides
than mdN. The active site residues Leu45(47), Tyr45(47) and Leu102(104)
of human and murine cdN (the residue numbers in parenthesis refer to murine cdN) form a more favorable binding surface for purine nucleotides than
the corresponding Ile133, Trp76 and Trp96 of mdN. Fig. 15 shows the mdNdGMP structure (Paper I) and the murine cdN-dGMP structure (Paper II)
superimposed.
dGMP fits nicely in the active site of cdN, with favorable distances to
catalytic residues.
In mdN, dGMP binds slightly displaced from catalytic residues. Its phosphate moiety is displaced from Asp43 (second aspartate of motif I), the residue that is proposed to donate a proton to the leaving nucleoside during the
catalysis (Fig. 11). This longer distance probably makes the transfer of a
proton from Asp43 to the nucleoside less efficient, causing slower catalysis.
This, in combination with lower affinity, probably causes the low activity of
mdN towards purine nucleotides.
Fig. 15. Superposition of the mdN-dGMP structure (light) and the murine cdNdGMP structure (dark).
35
Nucleoside analogue recognition in mdN
The D41N variant of mdN was also used to study recognition of the monophosphorylated NAs d4TMP, AZTMP and BVdUMP (Paper I and Paper
II). The chemical structures of these NAs are shown in Fig. 7, and their relative activities with cdN, mdN and cN-II are shown in Table 5.
The anti-HIV analogue d4TMP binds nearly identically to dTMP, with
the exception of the missing 3’OH (Fig. 16). The low activity for d4TMP
indicates the importance of the 3’OH for retaining enzyme activity.
There is a tight interaction of the 3’OH of dTMP with the backbone of
mdN. In all solved structures of 5’-(d)NMP complexes of both mdN and
cdN, the 3’OH forms tight interactions with the protein (however, sometimes
the 3’OH forms a hydrogen bond with Asp43 (mdN) instead of the main
chain). This indicates that most analogues with substituents at the 3’-position
would be poor substrates for mdN and cdN.
Fig. 16. Superposition of the mdN-d4TMP structure (dark) and the mdN-dTMP
structure (light).
The mdN-AZTMP structure shows another example of how an unfavorable binding in the pocket for the 3’OH, negatively affects catalytic efficency. The low activity towards AZTMP could be explained by the interactions of the azido group with the protein (Fig. 17). The azido group at the 3’position of AZTMP is too large to fit in the pocket for the 3’OH. This
probably greatly reduces substrate affinity.
36
Fig. 17. Superposition of mdN-AZT-MP (dark) and mdN-dTMP (light), the azido
group is marked AZ.
The analogue BVdUMP is a significantly better substrate for mdN than
AZTMP and d4TMP, probably because BVdUMP is substituted on the base
instead of the 3’-position. It fits relatively well in the active site of mdN,
although the base is turned 180 degrees relative the other substrates.
However, BVdUMP is a very poor substrate for cdN. The reason for this
might be that Arg47 and Tyr65 of human cdN possibly would clash with the
bromovinyl group of BVdUMP (Fig 18).
Fig. 18. Superposition of mdN-BVdUMP and human cdN. Residues R47 and Y65
belong to cdN, S78 and W96 to mdN. Clashes indicated by dotted lines.
37
Implications for the catalytic mechanism
The substrate complexes of mdN and cdN mimic the reaction intermediate 1
(Fig. 11). Intermediate 2 is supported by the structure of human cdN in complex with deoxyuridine and AlF4- (Paper II). Fig. 19 shows structures that
mimic intermediates 1 and 2. Although the deoxyuridine-AlF4- complex is
hexacoordinated, it still to some extent mimics the pentavalent substrate
intermediate 2. The deoxyuridine-AlF4- complex has a somewhat similar
charge distribution and geometry as the hypothetical pentavalant substrate
intermediate.
Fig. 19. Superposition of human cdN-AlF4--deoxyuridine (light) and mdN (D41N
variant)-3’dTMP (dark) structures.
38
Structure of human cN-II (Paper III and IV)
One of the aims in studying cN-II by x-ray crystallography was to better
understand the substrate specificity of this enzyme, which could increase the
understanding of cN-II’s role in NA resistance.
cN-II is allosterically regulated and probably has two effector sites, of
which one was well characterized (Paper IV). Important aspects of its
mechanism of regulation were also elucidated in Paper IV, see below.
Characterization of 2 effector sites
cN-II was initially crystallized in ~2 M of MgSO2. The sulfates efficiently
bound at several phosphate sites in the protein, probably stabilizing its structure. As a consequence, we could not bind nucleotides (e.g. effectors) or
other phosphate ligands to cN-II in the crystal, since the phosphate sites were
already occupied by sulfate.
However, we could compete out sulfate from the active site by binding
BeF3- covalently to catalytic Asp52 (first aspartate of motif I). We could also
bind adenosine to the protein, which bound with high occupancy to a site
that we named effector site 1. We also found less defined density at another
location that we interpreted as the adenine base of adenosine. We modeled in
adenosine in this site that we named effector site 2 (Paper III). We could
not confirm this site in later studies (Paper IV). However, a positive patch
(Q420RRIKK) in this site that bound two sulfate ions indicates this being
phosphate binding sites. This supports the notion that effector site 2 is a
binding site for nucleotides (Fig. 20). Possibly, a structural change is needed
to complete this effector site.
The structure shows cN-II as being a tetramer of two dimers (Fig. 21),
which is consistent with biochemical studies indicating that it functions as a
tetramer.
39
Fig. 20. cN-II in complex with two adenosines. AS=Active site. ES1=Effector site 1.
ES2=Effector site 2. Adenosines and sulfates are shown as sticks.
Activator recognition in effector site 1
The D52N variant of cN-II was co-crystallized with activator bound to effector site 1 and substrate and Mg2+ bound in the active site. Effector site 1 is
located close to the subunit interface between the dimers of the tetramer. In
this interface, the phosphate moieties of activators bind (Fig. 21).
In the case of dATP and ATP, a Mg2+ is coordinated between the phosphate moieties of the two nucleotides (Fig. 22). This Mg2+ neutralizes two
negative charges from the activators. Because of the well-defined electron
density and symmetric binding of dATP and ATP, these are likely to be
“true” effectors for this site (Fig. 22). Mg2+ binds at the 2–fold axis between
the dimers of the tetramer.
40
Fig. 21. Tetrameric structure of cN-II, with activator bound in effector site 1 and
substrate bound in the active site, shown as spheres.
Fig. 22. dATP bound in effector site 1. dATP* belongs to the adjacent subunit.
In contrast, the activator Ap4A binds with its phosphates in two alternative conformations. The activator 2,3-BPG also binds in two alternative conformations in the dimer interface. It traverses the 2-fold axis between the
subunits.
41
Mechanism for activation of cN-II
One of the first very important observations that we made when solving
structures of the D52N variant of cN-II, was that neither substrate nor Mg2+
bound to the active site of cN-II without an activator bound to effector site 1.
After soaking the crystals in reservoir solution with 10 mM of substrate and
10 mM of Mg2+, the structure obtained was still of the apoprotein. This suggests that the mechanism of activation of cN-II functions through a drastic
increase in substrate affinity.
Extensive kinetic data is available on how activators affect Km and Vmax
of cN-II (43-45, 47, 52), but the different studies are contradictory and it is
difficult to derive a consensus from them.
Table 6 shows how various effectors affect the activity of cN-II purified
from human placenta.
Table 6. Activity of cN-II, with/without activator and Pi
Activator
Reaction velocity (nmole/min)
None
0.6
dATP
26.9
ATP
20.5
GTP
12.0
2,3-BPG
17.8
Pi
0.3
ATP + 1 mM Pi
11.1
1.2
ATP +4 mM Pi
100 μM IMP was used as substrate, the activators were at 3 mM (44)
42
By comparing the apoprotein with activator/activator+substrate complexes, we could see significant changes in the structure that enabled us to
elucidate a mechanism for the allosteric activation of cN-II.
An α−helix seen in the complex structures is disordered in the apoprotein.
We have called this helix, helix A. (Fig. 23).
Fig. 23. IMP and Mg2+ bind in the active site and 2,3-BPG binds in effector site 1.
The apoprotein lacks helix A.
Helix A contains Asp356 of motif IV, a residue that is very conserved between HAD superfamily enzymes. Substitution of this aspartate into an asparagines or serine abolished the activity in mdN, PSP, HAD, and SERCA
(142-144 and unpublished data on mdN). The role of this residue is still
unknown.
43
The hydrogen bond pattern in the Mg2+-phosphate site is destroyed in the
apoprotein, compared to the complex structures (Fig. 24). Upon activator
recognition, structural changes occur that stabilize the hydrogen bond pattern
in the active site: Phe354 turns from the active site and Asp356 enters instead. There is also a conformational change in the conserved Lys292 of
motif III.
The activator stabilizes helix A by forming a hydrogen bond to Lys362
and by π−stacking with Phe354. The mechanism of activation of cN-II, is
probably initiated by the binding of an activator in effector site 1. This induces the formation of helix A containing Asp356 that completes the Mg2+phosphate binding site, whereafter the Mg2+ and substrate can bind.
The mechanism of activation through effector site 1 is, to our knowledge,
a novel type of mechanism. It might be that this mechanism is conserved in
other HAD superfamily enzymes that are regulated such as the P-type ATPases.
Fig. 24. Mechanism of activation. Left: Active site of apoprotein. Right: Activator
Ap4A bound in effector site 1 and GMP bound in active site.
44
Covalent modification on Asn52
Recently we solved a structure of the D52N variant of cN-II with a very
surprising covalent modification on Asn52 (corresponding to the first Asp of
motif I) (Fig. 14). We obtained this modification only when soaking crystals
in reservoir solution and activator, without Mg2+. We modeled this modification as a carboxylation (Fig. 25).
Fig. 25. Covalent modification of Asn52 modeled as a carboxy-Asn. Dotted lines
from Lys292 indicate hydrogen bonds, whereas the dotted line from Asp356 indicates the short distance (3.6 A) to the β-C of Asn52. a) Fo-Fc map calculated at 4σ.
b) Superposition of cN-II-carboxy-Asp structure and cN-II-GMP-Ap4A structure,
with helix A shown.
Having observed this modification, we speculated on the chemical background for its formation. Candidates as reactants are bicine and OH- from the
crystallization solution. One possibility might be that the mechanism for the
formation of this carboxylation involves a nucleophilic attack by Asn52 on
bicine.
Since asparagine is a poor nucleophile, Asn52 probably has to be highly
activated, in order to become sufficiently nucleophilic. One speculative
chemical route for this to happen is an enolation of Asn52, by the removal of
a proton from the β-C. If this is the case, then something within close proximity of the β-C of Asn52 has to be the activating agent. An important observation is that there is no room for a water molecule or OH- close to the βC of Asn52. The closest candidate for activating Asn52, is Asp356 (second
Asp of motif IV). Could Asp356 abstract a proton from the β-C of Asn52?
45
Asp52 of the wild type enzyme is clearly a stronger nucleophile than asparagine. However, it is not clear if Asp52 needs to be activated or not in
order to perform the nucleophilic attack on the substrate. By coordinating the
Mg2+, it temporarly loses some of its nucleophilicity, which could be a reason for the need of activation. It might, however, be possible that the substrate phosphate moiety (which also forms an electrostatic interaction with
the Mg2+) replaces Asp52 in neutralizing a charge of the Mg2+. This would
free the negative charge of the Asp to preform the nucleophilic attack, without activation.
If Asp52 needs to be activated to perform the nucleophilic attack, this
might occur through an enolation (similar as described above) (Fig. 26). The
enolation would generate two single bonded C-O moieties where one coordinates the metal and the other attacks the substrate phosphate.
Fig. 26. Hypothetical activation of D52 by an enole reaction.
46
Substrate recognition in cN-II
Several structures of the D52N variant of cN-II were solved in complex with
various substrates together with various activators (Table 4). The substrates
IMP, GMP, dGMP and UMP were bound to the active site of cN-II together
with activator bound in effector site 1. The suboptimal substrate UMP bound
with somewhat less defined density than the other substrates. AMP could not
be bound to the protein in the crystal.
From the recognition of IMP/GMP/dGMP it becomes clear how cN-II
discards AMP as substrate: Arg202 and Asp206 interact favorably with the
6-carbonyl group and NH1 group of IMP/GMP/dGMP, respectively (Fig. 1
and 27a). These two residues probably repel the amino group and N1 group
of AMP, respectively.
From the recognition of UMP, we can similarly explain how cN-II discards CMP as substrate: Arg202 that favorably binds the 4-carbonyl group of
UMP, would repel the amino group of CMP.
Probable explanations for why cN-II generally prefers purine over
pyrimidine nucleotides are (1) the fewer interactions that pyrimidine nucleotides have with the protein compared to purine nucleotides, and (2) that the
2-carbonyl group (that is present in all pyrimidine nucleotides) binds unfavorably close to Phe157 (Fig. 1 and 27b).
Lys215 might provide the preference of cN-II for ribonucleotides over
deoxynucleotides, since it creates a hydrophilic environment suitable for a
2’OH rather than a hydrophobic C2’.
Fig. 27. Substrate recognition in cN-II. a) IMP b) UMP.
47
The above observations strongly suggest that cN-II has a very poor affinity for cytosine and adenine based NAs. This further suggests that cN-II does
not directly contribute to resistance towards araC, dFdC, CdA and other
commonly used cytidine and adenine based NAs in treatment of cancer.
Instead guanine based NAs, such as ribavirin (Fig. 7) are likely to be directly affected by cN-II. Ribavirin was shown to be phosphorylated by cN-II
at a higher rate than by the dNKs (132).
48
Structure of human cN-III
The structure of cN-III was solved with molecular replacement using the
92% sequence identical murine cN-III structure (pdb code 2BDU) as a template. It was solved as apoprotein (2.7 Å), in complex with beryllium
trifluoride (BeF3-) (2.5 Å) and in complex with Pi (3.0 Å). Both the BeF3complex and the Pi complex has Mg2+ bound in the active site.
The numbering of the amino acids in the structure corresponds to isoform
2 (Q9H0P0-2), since the protein was expressed using cDNA of this isoform.
In order to get crystals, a construct was made that lacks the first 13 amino
acids. All residues expressed (Asn14–Leu286) are well defined in the structure.
Mapping of cN-III deficiency substitutions
cN-III is thought to catabolize UMP and CMP originating from RNA in
erythrocytes. Mutations in the cN-III gene have been found in patients with
hemolytic anemia. Several of these give amino acid changes: D87V, L131P,
N179S, G230R, C63R, Q143del, G157R and I247T.
It was shown that the L131P substitution decreased the stability of the enzyme (69). The G230R substitution gave lower affinity for CMP, suggesting
that Gly230 is important for substrate binding (69). The D87V, N179S,
G157R and I247T substitutions strongly reduced enzyme activity (67, 68,
70), whereas the L131P and C63R substitutions gave only moderate alterations in activity (70). The Q143 deletion did not significantly change enzyme
activity although it reduced the stability (66). The G157R variant was very
unstable (67).
The cN-III structure enabled mapping of the above-mentioned substitutions, providing structural evidence for how they can affect protein function
and stability. Four of the substitutions were already discussed based on the
murine cN-III structure (71), reaching conclusions to some extent similar to
ours. Fig. 28 shows Asp87, L131, N179 and G230, and also those residues
that we predict can participate in recognizing the base moiety of the substrate. Fig. 29 shows the location of residues C63, Q143, G157 and I247.
Implications based on our human cN-III structure:
The D87V substitution might lead to a hydrophobic collapse in the cap
domain, that probably affects residues that take part in recognition of the
base moiety of the substrate.
The L131P substitution probably changes the secondary structure of adjacent residues. Since Leu131 is close in space to motif I, it might affect the
structure of the Mg2+-phosphate site.
Asn179 forms tight hydrogen bonds to three main chain carbonyl groups
(Ala154, Gly155 and Ile197) and one main chain amide (Ile197). These resi49
dues are close in space both to catalytic residues and residues that possibly
participate in base recognition. The structure around the N179S substitution
is probably destabilized, affecting residues of the active site.
The G230R substitution could affect the fold of the protein, since an arginine at this position would clash into the adjacent secondary structure element (Val199 or Phe200). This could possibly affect the ability of the cap
domain to close over the core domain, thus reducing the substrate affinity.
Another possibility is that the arginine adapts its conformation so that it fills
up the active site and in that way inhibits substrate binding.
The C63R substitution might affect the structure of the cap domain, but is
rather far from the active site, which correlates with its low effect on enzyme
activity.
The Q143 deletion is also located rather far from the active site, which
correlates with its low effect on enzyme activity.
The G157 residue constitutes the tight turn between strand S2 and the following helix. Glycine is often positioned at tight turns in proteins since it has
no side chain and is therefore not as sterically restricted as other residues.
The G157R substitution probably strongly affects the tertiary structure,
which could explain the low stability of this variant.
The I247T substitution is located on strand S5 in the hydrophobic core of
the core domain close to motif IV, which probably affects the structure of
the catalytic phosphate site, affecting catalysis.
50
Fig. 28. Human cN-III-BeF3-Mg structure. Residues that are substituted are shown
as spheres, and residues of the cap domain (light) that might be involved in substrate
recognition are shown as sticks. BeF3-, covalently bound to Asp38 (sticks) and Mg2+
(sphere) are also shown.
51
Fig. 29. Human cN-III-BeF3-Mg structure. Residues that are substituted or deleted
are shown as spheres. Residues D227 and D231 of motif IV and BeF3-, covalently
bound to Asp38 are shown as sticks. Mg2+ is shown as a sphere.
52
Phosphotransferase activity of cN-II and cN-III
The role of the Thr/Val of motif I has yet not been biochemically investigated. We speculate that a Thr at this position might have a role in providing
5’NTs with phosphotransferase activity. Of the intracellular 5’NTs, only cNII and cN-III have a Thr at this position (Table 4). These two are also the
only 5’NTs that possess phosphotransferase activity.
This Thr appears to destabilize the conformation of the first Asp of motif
I. In structures of both cN-II and cN-III, we observe that this Asp can flip
and form a hydrogen bond to the Thr. This would not be possible with the
hydrophobic Val. Fig. 30 shows cN-III in complex with Pi where it has this
Asp in a flipped conformation, compared to its conformation with BeF3bound covalently.
Fig. 30. Superposition of the cN-III-BeF3 (light) and cN-III-PO4 (dark) structures.
53
Future perspectives
Structural insights into the substrate recognition of cytosolic 5’NTs could
assist efforts directed towards the understanding of 5’NTs involvement in
NA resistance. It could direct the choice of which 5’NT that should be used
for studying a specific NA. For instance, our structures of cN-II suggest that
it does not take adenine or cytosine based nucleotides as substrates. This
indicates that cN-II is not directly involved in resistance to treatment with
cytosine and adenine based NAs.
Characterization of the substrate recognition and implications for the transition states of 5’NT’s could greatly assist in efforts directed at generating
inhibitors of 5’NTs. The correct combination of NA with an inhibitor of a
5’NT might reduce NA resistance.
We are currently aiming at determining structures of substrate complexes
of cN-III, since the structural basis for its specificity is still unknown. We
would also like to solve the structures of cN-IA and cN-IB, to complete the
structural view of human intracellular 5’NTs. The structural basis for their
substrate specificity would provide a more complete understanding of intracellular 5’NT specificity. Although these three 5’NTs appear to have tissuespecific expression, they might be relevant for development of drugs that are
directed toward those tissues.
The complicated regulation of cN-II is yet not fully understood. It would
be of great interest to solve the structure of the full-length construct to analyze the possible effector site 2 and the role of its C-terminal acidic stretch.
Structures of cN-IA and cN-IB would also show the structural basis for their
allosteric regulation.
54
Acknowledgments
I would like to express my gratitude to everyone that has encouraged me
during my PhD studies, which have been a very exciting journey!
Especially I would like to thank:
Pär, for accepting me as a PhD student and for your encouragement, good
supervision and trust! I also thank you for providing a very inspiring atmosphere in the lab!
My collaborators in Padova: Vera Bianchi, for your great support in the
process of writing papers and for scientific guidance! Benedetta Ruzzenente
and Chiara Rampazzo for the purification of mdN and cdN, and for nice
discussions and nice company during meetings.
Agnes, for your support, collaboration, friendship and fun company during many travel adventures!
Stefan Nordlund for being a good boss! I would also like to thank you for
sending me to Zurich as an Erasmus student, and you and Peter Brzezinski
for recommending me to ”Forskarskolan”, since those opportunities helped
me a great deal!
Pål, Susanne, Tomas and Urszula of Team 1 at SGC, for a nice collaboration on the wild type cN-II, cN-III and UCK projects.
Karl-Magnus for always being very helpful and supportive! Daniel G for
being a great labasse-kompis! Audur, Sue-Li, Lola and Anna-Karin for
friendship and for providing some balance against the boys, Monica V for
friendship and all the fun during travels ☺, Damian, Daniel MM, Tobias,
Ulrika, Amin, Marie, Marina, Pelle, Maria, Hanna, Said, Anna, Henrik,
Heidi, Albert, Martin Hö, the newcomers Christine, Christian and Cedric, all
other past and present members of the PN-group, the SGC people and the
administrative staff at both SU and KI (especially Elisabeth) for being very
nice and helpful!
Jag skulle också vilja tacka min kära familj, mina underbara utanförlabbet-vänner och min gosiga pojkvän David för det stöd och tålamod som
jag fått från er! TACK!
55
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