Biochem. J. (2009) 417, 401–409 (Printed in Great Britain)
401
doi:10.1042/BJ20081986
REVIEW ARTICLE
Evolution of protein phosphatases in plants and animals
Greg B. G. MOORHEAD1 , Veerle DE WEVER, George TEMPLETON and David KERK
Department of Biological Sciences, University of Calgary, 2500 University Dr., N.W. Calgary, Alberta, Canada T2N 1N4
Protein phosphorylation appears to be a universal mechanism
of protein regulation. Genomics has provided the means to compile inventories of protein phosphatases across a wide selection of
organisms and this has supplied insights into the evolution of this
group of enzymes. Protein phosphatases evolved independently
several times yielding the groups we observe today. Starting
from a core catalytic domain, phosphatases evolved by a series
of gene duplication events and by adopting the use of regulatory
subunits and/or fusion with novel functional modules or domains.
Recent analyses also suggest that the serine/threonine specific
enzymes are more ancient than the PTPs (protein tyrosine
phosphatases). It is likely that the latter played a key role at the
onset of metazoan evolution in conjunction with the tremendous
expansion of tyrosine kinases and PTPs at this point. In the
present review, we discuss the evolution of the PTPs, the serine/
threonine specific PPP (phosphoprotein phosphatase) and PPM
(metallo-dependent protein phosphatase) families and the more
recently discovered phosphatases that utilize an aspartate-based
catalytic mechanism. We will also highlight examples of
convergent evolution and several phosphatases which are unique
to plants.
INTRODUCTION
and through biochemistry, molecular biology and most recently
genomics, we can now catalogue the phosphatase complement
of organisms and begin to examine evolutionary relationships
between the protein phosphatases. This review will discuss the
spectrum of protein phosphatases, evolutionary relationships,
convergent evolution and the role of phosphatases in the evolution
of metazoans.
There are few individuals whose contribution to science has had a
more profound effect on the study of biology than that of Charles
Darwin. It is an honour to write an article on protein phosphatase
evolution to commemorate his 200th birthday and 150 years since
the 1859 publication of “On the Origin of Species”. We live
in a time when we are beginning to understand evolution at a
molecular level and, having passed into the post-genomic era,
we are currently seeing a transition to an age of evolutionary
and comparative genomics, which ideally will culminate into
evolutionary proteomics. These emerging fields will undoubtedly
have a profound effect on not only evolutionary and genomic
biology, but on biological research in general.
Recent studies have highlighted the vast number of modified
proteins within proteomes and the often large number of
functional groups found on any given protein. Many studies
have also reinforced the view that protein phosphorylation is a
common means to the regulation of most cellular processes, is
ancient in origin, and is probably a universal covalent modification
among organisms [1–3]. This is highlighted by the fact that
protein kinases and phosphatases, which respectively add and
remove phosphate on proteins, constitute 2– 4 % of the genes
in a typical eukaryotic genome [4,5]. Early studies on protein
phosphorylation focused on its role in modifying enzyme activity,
while more recently it has been found to play a role in protein
localization, protein turnover and to provide a specific interaction
site for other proteins [6]. Like protein kinases, the study of
protein phosphatases began in the field of glycogen metabolism
Key words: genomics, kelch phosphatase, laforin, phylogenetics,
protein tyrosine phosphatase (PTP), starch excess 4 (SEX4),
TFIIF associating component of CTD phosphatase/small CTD
phosphatase (FCP1/SCP).
PROTEIN PHOSPHATASE CLASSIFICATION
Proteins can be phosphorylated on nine amino acids (tyrosine,
serine, threonine, cysteine, arginine, lysine, aspartate, glutamate
and histidine) with serine, threonine and tyrosine phosphorylation
being predominant in eukaryotic cells and playing key regulatory
roles [5]. Although phosphorylation on aspartate and histidine
residues in bacterial two-component systems have been well
characterized, several recent phospho-proteomic studies have
shown bacterial protein phosphorylation to be more predominant
than originally thought [7–9]. In three bacterial species, 63–
79 proteins were found to be phosphorylated, with a surprising
distribution on serine, threonine and tyrosine of approx. 69, 22
and 9 % respectively. In humans, phosphorylation on serine,
threonine and tyrosine is approx. 86.4, 11.8 and 1.8 % respectively
[10]. The enzymes that dephosphorylate these amino acids
(serine, threonine and tyrosine) form four groups based on
unique catalytic signatures/domain sequences and substrate preference. The majority of phospho-serine and phospho-threonine
dephosphorylation is accounted for by the PPP (phosphoprotein
Abbreviations used: CDC, cell division cycle; CTD, C-terminal domain; DSP (or DUSP), dual-specificity phosphatase; EPM2A, epilepsy of progressive
myoclonus type 2 gene A; EYA, eyes absent; FCP/SCP, TFIIF (transcription initiation factor IIF)-associating component of CTD phosphatase/small CTD
phosphatase; HAD, haloacid dehalogenase; LMPTP, low-molecular-mass PTP; MAPK, mitogen-activated protein kinase; MAPKP, MAPK phosphatase;
MTM, myotubularin; PAS2, Pasticcino 2; PP1 etc., protein phosphatase 1 etc.; PPKL, protein phosphatase 1 and kelch-like; PPM, metallo-dependent
protein phosphatase; PPP, phosphoprotein phosphatase; PRL, phosphatase of regenerating liver; PTB domain, phosphotyrosine-binding domain; PTEN,
phosphatase and tensin homologue deleted on chromosome 10; PTP, protein tyrosine phosphatase; PTPLA/B, PTP-like (proline instead of catalytic
arginine), member A/B; RLK, receptor-like kinase; RPTP, receptor PTP; SEX4, starch excess 4; SH2 domain, Src homology 2 domain; SIX, Sine oculis ;
TFIIF, transcription initiation factor IIF; UBLCP1, ubiquitin-like domain containing CTD phosphatase 1.
1
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2009 Biochemical Society
402
G. B. G. Moorhead and others
phosphatase) [PP (protein phosphatase) 1, PP2A, PP2B (or
PP3), PP4, PP5, PP6 and PP7] and PPM (metallo-dependent
protein phosphatase; PP2C) families and includes the enzymes
shown in parentheses. Remarkably, the PPM and PPP families
are unrelated in sequence and are probably evolved from two
unique ancestral genes, but converged to have highly related
structures at the catalytic centre [11]. The PTP (protein tyrosine
phosphatase) family is defined by the catalytic signature CX5 R
and is very diverse in domain structure and substrate preference
with some family enzymes now recognized to dephosphorylate
complex carbohydrates, mRNA and phosphoinositides [12,13].
Finally, for the most recently classified group, the aspartatebased phosphatases, catalysis is driven by an aspartic acid
signature (DXDXT/V) and includes the FCP/SCP [TFIIF
(transcription initiation factor IIF)-associating component of CTD
(C-terminal domain) phosphatase/small CTD phosphatase] and
HAD (haloacid dehalogenase) family enzymes. We refer readers
to several excellent reviews on phosphatase classification and
functions [12–19].
SPECIFICITY THROUGH MODULAR DOMAINS AND REGULATORY
SUBUNITS
Having a complete inventory of phosphatases for several
organisms reveals many aspects of this group of enzymes, with one
striking feature being immediately apparent: ancient phosphatase
domains functionally evolved by docking to novel regulatory
subunits (most PPP members), or through additional domains
that were acquired by fusing with these catalytic domains at the
gene level (PTP, PPM and aspartate-based families) (see Figure 1).
Although the PTP core domain has also evolved to accommodate
specific substrates, other phosphatase catalytic subunits typically
display broad substrate specificity in vitro. As a result, specificity
of function is brought about by an array of modules or domains
and regulatory subunits. Domains and subunits provide a means of
subcellular targeting, allow docking of other proteins and inositol
lipids and often permit regulation by covalent modifications [20].
The evolutionary strategy of adding domains on to a core unit was
first noted for the PTPs, but is also apparent for the PPM (PP2C)
family (especially in plants) and a number of other phosphatases
(e.g. PP5). The PPP family members PP1 and PP2A are some
of the most highly conserved enzymes known. Conversely, they
obtain specificity for their catalytic engines via interactions with
many novel regulatory subunits. The best example of this is PP1,
with nearly 100 recognized regulatory subunits (Figure 1). Two of
the more recently identified members of the PPP family, PP4 and
PP6, followed a similar evolutionary route as PP1 and PP2A,
with a remarkable conservation of regulatory subunits across
species [21,22]. For instance, the PP4-interacting proteins, R2
and R3α and β of humans, have readily apparent orthologues in
the Saccharomyces cerevisiae genome that have been shown to
complex with the yeast PP4 catalytic subunit [23].
PTPs
The PTPs are defined through their common signature motif
(CX5 R) that drives catalysis [13]. It is thought that this motif and
catalytic mechanism evolved independently three times to yield
three groups or classes of PTPs [12]. The human genome encodes
more than 100 PTPs with orthologues for all but one in mouse
[12]. Of the more than 100 PTPs, 11 are catalytically inactive
and 16 dephosphorylate either glycogen (1), mRNA (2) or
phosphoinositides (13), and not proteins. It is thought that the
remainder of these PTPs dephosphorylate phosphotyrosine and,
c The Authors Journal compilation c 2009 Biochemical Society
Figure 1
Distinct evolutionary routes for protein phosphatases
Catalytic and regulatory modules have evolved by two means to accommodate specific
substrates and/or functions. (A) represents evolution by domain fusion on a genome level,
exemplified by PTP evolution. This evolutionary mode is also followed by some PPP members
(e.g. kelch-like phosphatases and aspartate-based phosphatases). The ancestral catalytic
domain (blue rectangle) fused with a regulatory module (red oval). Subsequent evolution of
regulatory modules (pentagon, oval, triangle) confer unique properties to the catalytic domain.
Modifications in the catalytic domain throughout evolution render further substrate specificity
(depicted by the shading of the catalytic domain). Some catalytic domains have become inactive
(red cross). (B) reflects evolution via complex formation on a proteome level, conceptualized
using PP1 as an example. The catalytic subunit (green oval) accommodates novel substrates via
interaction with numerous regulatory subunits (rounded triangle, pentagon, star). The RVxF/W
motif (black bar) is the primary PP1-docking motif.
in some cases, also phosphoserine and phosphothreonine. Class I,
which include the classic and dual-specificity enzymes, have a
common PTP domain structural fold and are by far the largest
group of PTPs and are further divided into subfamilies. The
classic enzymes (receptor and non-receptor) are given this name
as they defined the PTPs and they all dephosphorylate tyrosine
residues. The DSPs (dual-specificity phosphatases; also known
as DUSPs; this group includes all non-protein phosphatase PTPs)
are further divided into the MAPKP [MAPK (mitogen-activated
protein kinase) phosphatase], slingshot, PRL (phosphatase of
regenerating liver), atypical DSP, CDC14 (cell division cycle
14), PTEN (phosphatase and tensin homologue deleted on
chromosome 10) and MTMs (myotubularins). Class II and
III are represented by the LMPTP (low-molecular-mass PTP)
and Cdc25 isoforms and are thought to have evolved from a
bacterial rhodanese-like enzyme and a bacterial arsenate reductase
respectively (see [12,13,24] for further details).
In plants, no true protein tyrosine kinases exist, even though in
Arabidopsis there are at least 1055 protein kinases (compared with
518 in humans) of which ∼ 630 are RLKs (receptor-like kinases).
These RLKs are most like human Pelle kinases that display
serine/threonine kinase activity, and as such it is believed that
all plant RLKs are serine/threonine kinases [25,26]. Consistent
with the absence of tyrosine kinases is the near total lack of
plant phosphotyrosine-specific phosphatases in the PTP family
(i.e. classic enzymes) [27–29]. As with plants, other genomes
examined that were found to lack tyrosine kinases also have very
few PTPs [30]. For example, most species that belong to the
apicomplexa (protozoans) have no class II, class III or classic
PTPs and only a very few DSPs (in this case, PRL and MAPKP).
Apicomplexa genomes also have no obvious protein tyrosine
kinases [30]. As will be discussed below, it is now thought that
the PTPs that dephosphorylate phosphotyrosine evolved before
Protein phosphatase evolution
tyrosine kinases due to the action of promiscuous serine/threonine
kinases that phosphorylate tyrosine residues. The appearance of
true tyrosine kinases coincides with a dramatic expansion of the
PTP superfamily just before the evolution of metazoans.
Sequence and structural analysis of the class I PTPs reveals a
universal structural fold supporting the concept of evolution from
a common ancestral gene [12]. Consistent with this is evidence
of gene duplication. The highly sequence-related pairs TCPTP
(T-cell PTP)/PTP1B, PTPα/PTPε and PTPD1/PTPD2 each share
very similar exon structures in the genomes of human, mouse
and chimpanzee [31]. Human DUSP24 is localized adjacent to
DUSPs 13a and 13b indicating two ancestral gene duplications.
Humans have 21 RPTPs (receptor PTPs) and 12 of them have
a tandem arrangement of intracellular catalytic domains (D1 is
membrane proximal and D2 is membrane distal). With only one
exception, all D1 are active and all D2 are inactive, yet all D2s are
still well conserved compared with other PTP domains [13,32].
Phylogenetic analysis of PTP domains shows that D1 and D2
are from separate clades (i.e. all D1s are most like each other),
but group together when compared with all other PTP domains,
indicating that the PTP domain duplicated within the ancestral
PTP gene before the whole gene duplicated and ultimately gave
rise to the 12 RPTPs [32]. Recent analysis has shown that
the catalytic cysteine residue of the CX5 R motif is susceptible
to oxidation due to the same properties that make it a good
nucleophile [33]. Nature has evolved two mechanisms to protect
this group from irreversible oxidation [34]. PTP1B has been
shown to form a reversible cyclic sulfenamide with the catalytic
cysteine residue forming a bond with the amide nitrogen of the
neighbouring serine residue. PTPs from the DSP, LMPTP and
CDC25 groups have been demonstrated to form disulfide bonds
between the catalytic cysteine residue and another cysteine residue
of the same protein [34]. Although the D2 domains of RPTPs are
inactive, they intriguingly maintain their CX5 R signature and
are inactivated by loss of other key catalytic amino acids. The
remarkable conservation of D2 PTP domains and specifically
the CX5 R motifs has been postulated to be due to the potential
role of these domains as cellular redox sensors [34].
Certain members of the PTP family exemplify how sequence
and function conservation can go hand in hand. The class I dualspecificity enzyme Cdc14 controls mitotic exit in metazoans and
yeast. Searching for this enzyme in plants and eukaryotic (green)
algae revealed a single orthologue in Chlamydomonas reinhardtii
with sequence coverage over both the N-terminal region and PTP
domain [27]. Cdc14-like protein(s) are also present in higherplant genomes, yet phylogenetic analysis showed that they form a
distinct clade, separate from characterized Cdc14 proteins. These
Cdc14-like sequences lack key catalytic and substrate targeting
residues and thus could not function as active phosphatases,
but seem to have been co-opted into protein–protein interaction
domains [27]. Although putative Cdc14 orthologues have been
noted in many protozoans, there is an absence of this PTP in all
apicomplexans, except Cryptosporidium [30].
The single class II enzyme, also known as LMPTP, is tyrosinespecific and widespread, being found in all kingdoms including
Archea and most bacterial species, and is one of only three PTPs
in yeast [12,35]. A survey of several bacterial LMPTPs shows that
the enzyme is 30– 40 % identical with the human enzyme. This
astonishing conservation suggests an ancient conserved function,
but to date no clear role has been defined for LMPTP.
The sole class III PTP, also known as Cdc25, dephosphorylates the cell-cycle protein kinase Cdc2 [36]. The Cdc25s display
sequence similarity to rhodaneses and are thought to have evolved
from a bacterial, rhodanese-like protein [12,27]. Our genomic
analysis of several plant and algal species for phosphatases related
403
to that of human Cdc25 revealed that, like Cdc14, there appears to be no Cdc25 in plants [27]. Several, but not all, unicellular
parasites have Cdc25 orthologues. A notable absence is again in
the apicomplexa [30]. Candidate plant and algal Cdc25s actually
form a clade with several arsenate reductases and not the wellcharacterized human and yeast Cdc25s suggesting that these
candidates are really arsenate reductases. These sequences also
lack the N-terminal regulatory domain found in true Cdc25s. This
insight, along with the lack of Cdc14 in plants, indicates that
the role that these phosphatases play in cell-cycle control was
established after ancestral plant and animals/yeast diverged, or
was lost early in the plant lineage.
Inactive PTPs
A number of enzymes are inactive, yet conserved, in many
organisms (e.g. the pseudokinases), which opens discussion
towards their biological function [37–39]. The phosphatases are
no exception to this, particularly the PTPs since the core catalytic
motif is well defined and inactive mutations are readily identified
[12,13]. Inactive PTPs display PTP-like domain structures, but
possess mutations in the cysteine or arginine residues of the
CX5 R motif, in an upstream conserved aspartate residue or in
combinations of the three, all exemplified by the STYX domain
proteins and the D2 domain of RPTPs [13,40]. In the case of
STYX, such alterations are thought to disrupt the enzymatic
activity of the phosphatase, yet potentially maintain interaction
with phosphorylated substrates ([12,40] and references therein),
not unlike how a 14-3-3 protein docks a phosphoserine or
threonine motif. Mutations must have occurred early in evolution
since various inactive PTPs {e.g. tensins, PTPLA/Bs [PTPlike (proline instead of catalytic arginine), member A/B]} are
conserved within eukaryotes [41,42].
The MTMs, a well-defined class I PTP subfamily, are phosphoinositide phosphatases specific for the 3-position of PtdIns3P and
PtdIns(3,5)P2 . These phosphoinositides provide docking sites for
proteins involved in endosomal–lysosomal membrane trafficking.
Only a few MTMs have been found in a number of apicomplexan
species, while we identified two presumably active MTMs in
Arabidopsis [27]. In humans, seven of the 16 MTMs are predicted
to be inactive [12]. Their characterization indicates that they play
essential roles, some of which are linked to human disease.
For example, loss of either MTMR2 or MTMR13, an active
and inactive MTM respectively, both cause a form of Charcot–
Marie tooth disease [43]. Hence, the inactive protein is equally
essential to prevent disease onset, and both proteins are thought to
interact.
Another family of potentially inactive PTPs are the tensins.
The tensin family has four members in humans (Tensins 1–
3 and Cten), each containing an SH2 (Src homology 2) and
PTB (phosphotyrosine-binding) domain at their C-termini [44].
Primary sequence analyses predict either an inactive PTP domain
[45] or a PTEN-C2-like domain in the N-terminal region of
Tensins 1–3 [46]. Cten lacks the N-terminal region and therefore
the putative PTP domain. Recently, Tensins 1 and 2 and Cten
expression were shown to stimulate cell migration whereas Tensin
3 expression had an inhibitory effect. Interestingly, our sequence
alignment shows that Tensin 3 maintains the CX5 R motif, whereas
Tensins 1 and 2 display mutations in these residues (cysteine to
asparagine and arginine to lysine respectively; results not shown),
whereas Cten obviously lacks the CX5 R motif.
Finally, the PTP-like proteins PTPLA/B are classified as
inactive PTPs, although they have no sequence homology with the
PTPs except for the CX5 R motif, where in their case the arginine
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G. B. G. Moorhead and others
residue is mutated into a proline [47]. Significant progress towards
the elucidation of their biological function has been made with the
PTPLA homologues in plants and baker’s yeast, PAS2 (Pasticcino
2) and Phs1 respectively. Reciprocal complementation studies
identified them as functional orthologues [48], while PTPLA/B
expression also rescues Phs1-impaired cells [49]. Both PAS2 and
Phs1 function as dehydratases in the elongase complex required
for the production of very-long-chain fatty acids. Further studies
are required to clarify the role of the CX5 R motif in these proteins
and the function of PTPLA in metazoan organisms.
Overall, these examples show that loss of key catalytic
residues, but overall domain conservation, led these proteins to
evolve from their initial function as phosphate-removing enzymes
towards different functions, e.g. phosphotyrosine-recognition
with potential scaffolding and localization capacities.
TYROSINE PHOSPHATASES AND METAZOAN EVOLUTION
Until recently, phosphotyrosine signalling was considered the
hallmark of intercellular communication of multicellular animals
as other unicellular organisms (including yeast) lack true tyrosine
kinases. Previously, a moderate selection of SH2 and PTP domains
were noted in the genomes of plants, fungi and dictyostelium
and in a few cases shown to be functional, as predicted [50 –
52]. Baker’s yeast, for example, has three PTPs [12,52,53] and
phospho-proteomics of several yeast species have shown a handful
of tyrosine phosphorylated proteins, even in the absence of true
tyrosine kinases [54–56]. This is explained by what has been
described as promiscuous serine/threonine kinase activity and
the action of the well-characterized MAPKK (MAPK kinase)
enzymes that phosphorylate MAPK activation loops in a TXY
motif [57].
The choanoflagellates are unicellular protists and, through
comparative genomics of mitochondrial DNA, they are currently
considered the closest living relatives of metazoans [58]. Recent
completion of the first choanoflagellate (Monosiga brevicollis)
genome has revealed an abundance of tyrosine kinases (∼ 128),
phosphotyrosine-specific PTPs (∼ 39) and proteins with the
phosphotyrosine-docking domains SH2 (∼ 123) and PTB (∼ 20)
[39,57,58]. Surprisingly, these numbers exceed that of any other
metazoan and support the concept that the phosphotyrosinesignalling machinery was present prior to the evolution of
multicellular animals.
These landmark genomic analyses tell us that although phosphorylation on tyrosine was present prior to metazoan evolution, it
had a limited role until tyrosine kinases evolved and with the tools
of kinase, phosphatase and phosphotyrosine-docking domain at
hand, a rapid expansion of all components occurred in some premetazoan ancestor that ultimately led to choanoflagellates and
multicellular animals. A comparison of the 38 classic (tyrosinespecific) PTPs of humans to the 39 M. brevicollis PTPs starkly
shows that only four are probably true orthologues, which presumably evolved in a pre-metazoan ancestor, while the remaining
independently evolved in each lineage with unique combinations
of signalling domains. This rapid expansion of the PTPs and
the phosphotyrosine-signalling system certainly played, to some
extent, a role in metazoan evolution. In contrast, as discussed
in [39,57], other well-recognized signalling domains/systems
(excluding small GTPases, phosphoserine/threonine kinases,
phosphatases and docking domains) are much more ancient and
appear in combinations that are common to both M. brevicollis,
metazoans and other eukaryotes, like plants and fungi, serving
as a reminder of how ancient other (non-PTP) phosphatases
are.
c The Authors Journal compilation c 2009 Biochemical Society
THE PPM FAMILY
The PPM family phosphatases, which include the PP2C
and pyruvate dehydrogenase phosphatase, are Mn2+ /Mg2+ dependent serine/threonine-specific enzymes that are resistant to
microcystin, okadaic acid and other classic toxin inhibitors of
the PPP family. Unlike most PPP members, the PP2C enzymes
do not have additional subunits but, like the PTPs, do display a
wide variety of additional domains that confer unique functions.
All eukaryotic PPM enzymes have 11 conserved motifs with nine
highly conserved amino acids, four of which are aspartate residues
that co-ordinate metal ions necessary for catalysis. Analysis of 16
archaeal genomes shows the presence of one PPM gene, while
bacteria display putative PPM genes in 27 out of 121 genomes
examined [59], grouped into two subfamilies. Interestingly,
subfamily II enzymes usually have additional domains with most
being GAF and PAS domains, while subfamily I genes are nearly
always lone catalytic domains [59]. While most bacterial species
with PPMs have one or a few PPMs, two Streptomyces
species have as many as 49 PPMs. Phylogenetic analysis of
these enzymes with eukaryotic PPMs reveals that the archaeal and
bacterial enzymes are rooted in the eukaryotic PPMs, suggesting
that the Mn2+ /Mg2+ -dependent enzymes originated in eukaryotes
and radiated into bacteria and archaea by horizontal gene transfer
[59]. Because PPM genes are spread widely throughout the bacterial kingdom, it is thought PPMs originated very early in the
eukaryotic lineage, then spread horizontally into bacteria early in
that lineage, and probably transferred several times. Support for
an early eukaryotic origin is upheld by the observation that PPM
phosphatases are found throughout eukaryotes, including plants,
and have no sequence homology with the ancient PPP enzymes.
The PPM enzymes are the largest phosphatase family in
plants, with 76 members in Arabidopsis thaliana. Phylogenetics
assembles them into ten subfamilies with six non-clustered genes
[60]. What is apparent from analysis of this group is the large
number of N- and C-terminal additions to these genes. They
are thought to confer specificity and/or localization to enzyme
function. These domains include MAPK-docking regions, FHA
(forkhead associated) domains, MORN (membrane occupation
and recognition nexus) repeats and putative transmembranespanning domains [60].
The PP2C members of humans have orthologues in most other
vertebrates [61]. Each vertebrate PP2C member (for instance
all PP2Cεs) clusters with the others, supporting the idea that
the family arose by a series of duplication events. A further
phylogenetic reconstruction of the metazoan PP2Cs reveals that
all enzymes cluster into two groups with one group having
unique sequences near the active site, as previously noted in
mammalian PP2Cs [61]. Analysis also indicates that two series
of rapid duplications took place, one before the emergence of
bilaterians and a second at the emergence of vertebrates. The
vertebrate-specific duplications have been linked to tissue-specific
expression patterns and may contribute to tissue development
in vertebrates. Interestingly, many other (non-phosphatase) gene
families, including the protein kinase subfamilies, evolved via a
series of duplication events [4], reminiscent of PP2C, supporting
the concept that expansion of signalling protein families was a
major driving force during key events of metazoan evolution.
THE PPP FAMILY
The PPP family is defined by three signature motifs (-GDXHG-,
-GDXVDRG- and -GNHE-) within a ∼ 280 amino acid catalytic
domain. PPP enzymes are regarded as very ancient, with most
members widely distributed in all eukaryotes and most bacterial
Protein phosphatase evolution
and archeal genomes having at least one PPP-like member. A
comparison of the PPP complements of humans and Arabidopsis
shows that plants have a complete absence of PP2B, but an
expansion of the number of PP1, PP2A, PP4 and PP6 genes and the
addition of a number of novel enzymes. Remarkably, the catalytic
domains of mammalian PP1s are 76 –88 % and 90 % identical
with plant and fungi PP1s respectively. Perhaps more remarkable
is that human PP1β is 100 % identical with rat, mouse, rabbit
and chicken and 97 % identical with the zebra fish, goldfish and
Atlantic salmon enzymes. Other noteworthy PPP complements
are found in the protozoan parasites Encephalitozoon cuniculi,
with only five PPP enzymes, and Entamoeba histolytica and
Trichomonas vaginalis both having large expansions with 81
and 169 predicted PPPs respectively [62], compared with 13 and
26 in humans and plants [27]. In addition there is the notable
absence of PP7 in yeast, but the presence of the unique PPP
members PPQ, PPZ1, PPZ2 and PPG1.
We have generated an unrooted tree for the PPP members
in humans and Arabidopsis thaliana (Figure 2), and the result
is consistent with similar analyses using a broader range of
eukaryotic PPP members [19,62,63]. PP5, PP7 and two plantspecific enzymes form a clade separate from other PPP members,
suggesting two distinct branches that probably separated early in
eukaryotic evolution. It has been proposed that the N-terminal
TPR (tetratricopeptide repeat) domains of PP5 and the Cterminal EF hand extensions of PP7 were originally separate
regulatory subunits and gene fusion yielded the products we
see today [62]. Because these extra domains are conserved
across eukaryotes, this too must have been an early event in
PP5 and PP7 evolution. With the presumed exception of the
Kelch-domain phosphatases, which were derived from a PP1
ancestor (Figure 1 and [62], discussed below), all other PPP
enzymes of this branch have regulatory subunits that define the
function of these proteins. As indicated previously, because a
majority of these additional regulatory subunits are present in
genomes of other eukaryotes and are often functionally conserved,
it is likely that most originated early in eukaryotic evolution
(Figure 1). This may explain why these PPP enzymes are so
highly conserved. Having these interaction partners that define the
function of the catalytic subunit certainly would hinder additional
evolution of the catalytic engine. For instance, the majority of the
PP1 regulatory subunits have a primary docking site called
the RVXF/W motif that allows interaction with PP1 in a
hydrophobic cleft on the surface of the enzyme [64–66].
Undoubtedly this docking site was exploited in the history of
PP1 allowing it to be recruited again and again to specifically
dephosphorylate new target proteins by incorporation of this short
docking motif in the regulatory subunit (Figure 1). Consistent
with this proposal is the knowledge that nearly 100 PP1-binding
proteins have now been identified and it is expected that there are
at least that many more several times over [14,24].
A unique PP1-related enzyme with extra domains is the PPKLs
(PP1 and kelch-like). The PPKL enzyme family is defined by
their large, kelch repeat containing N-terminal extensions and Cterminal PP1-like phosphatase domain. The kelch domain is a
widespread and ancient motif, with limited conserved residues
[67]. The founding members of the PPKL family are the
Plasmodium falciparium protein PfPPα [68] and four proteins
identified in A. thaliana [28,69]. BLAST searches with the kelchrepeat domain of these five proteins against translated genomes
identified further family members in the kingdom plantae (green
algae, vascular plants, mosses) and the superphylum alveolata,
with candidates in both the obligate parasitic apicomplexa and
the free-living aquatic ciliophora (Supplementary Figure S1
at http://www.BiochemJ.org/bj/417/bj4170401add.htm). We did
405
uncover PPKL members in Solanum lycopersicum (tomato) and
Vinis vinifera (grape) yet despite extensive searches, which
included newly sequenced genomes, we could not identify
orthologues in red algae, fungi or animals, nor in any noneukaryotic species, in keeping with previous reports [69,70].
Comparisons of the conserved residues in the PPP family
with the PPKL enzymes [71,72] indicate that most, but not
all, conserved PPP residues are maintained in this family (see
Supplementary Figure S2 at http://www.BiochemJ.org/bj/417/
bj4170401add.htm). This analysis further underscores how
(i) PPKLs contain five inserts in their phosphatase domain [68,69];
(ii) PPKL enzymes are more closely related to PP1 than to PP2A;
(iii) a significant number of PP1 family residues, particularly
those involved in small-molecule-inhibitor binding, either diverge
from the consensus or are preceded by peptide sequences which
may alter inhibitor binding [73]; and (iv) PPKL clades can be
defined by certain residues. For further details please refer to
Supplementary Figure S2 and [68–70].
The origin of the PPKL family is still under debate, as is
the origin of the apicomplexa, to which a large number of the
PPKLs are annotated (Supplementary Figure S1 and [74,75]).
We built a phylogenetic tree with the PPKL family members
using either domain separately (results not shown) or whole
protein sequences (Supplementary Figure S1; this Figure is
representative for each approach). We, and others, have found
PPKLs only in plants, green algae and several Alveolata
species [70]. This shared genotype between otherwise dissimilar
phyla support the current alveolate origin hypothesis whereby
a progenitor of (green and red) algae and plants yielded an
ancestral enzyme (i.e. the kelch domain/PP1-like enzyme), which
was retained in these organisms via secondary endosymbiosis
of the algae and subsequent nuclear gene transfer [76,77].
Further evolution of the alveolates and green plants could
thus produce the plant-related kelch-domain phosphatases. This
hypothesis could also explain why both apicomplexa and
green plants lack most PTP members. Finally, the current
PPKL distribution, which lacks members in the chromista, also
supports recent studies that question the strength of the current
grouping that yields the chromalveolate phylum [74,75]. We refer
readers to the following websites (http://tolweb.org/Eukaryotes/3
or www.discoverlife.org/mp/20m?tree = Life&flags = all) for
current classification schemes.
ASPARTATE-BASED CATALYSIS
While the other families of protein phosphatases have been
studied for more than 20 years, those utilizing aspartate-based
catalysis have only been noted in the last decade. The first
identified member of this family was FCP1, a TFIIF-associated
RNA polymerase II CTD phosphatase [78]. The aspartate-based
catalysis group is often divided into FCP/SCP-like and HADfamily phosphatases, but both actually share the same catalytic
motif and belong to different groups within the HAD superfamily
(http://www.ebi.ac.uk/interpro/). The HAD-family phosphatases
are ancient proteins, with FCP1 conserved throughout Eukaryota,
like members of the PPP family. However, unlike most members
of the PPP family, specificity of function has been gained through
the addition of other domains to the catalytic module, not
through association with other proteins (see Figure 1A for a
modular representation). One common additional domain is the
BRCT domain, which may function to direct these phosphatases
to phosphorylated substrates.
The evolutionary conservation of FCP1, as with other
phosphatases mentioned above, is completely logical where we
understand biochemical function. FCP1 conservation throughout
c The Authors Journal compilation c 2009 Biochemical Society
406
Figure 2
G. B. G. Moorhead and others
Phylogenetic analysis of the PPP family enzymes from Homo sapiens and A. thaliana
An unrooted tree was generated by comparison of catalytic domains from members of the PPP family from human and A. thaliana . Catalytic domains were aligned using ClustalX, hand-optimized
using GeneDoc, and the bootstrap tree was generated using the neighbour-joining function of ClustalX [27]. Proteins with an At suffix are from A. thaliana , while an Hs suffix denotes human
sequences. The Arabidopsis sequences are: TOPP1 (At2g29400), TOPP2 (At5g59160), TOPP3 (At1g64040), TOPP4 (At2g39840), TOPP5 (At3g46820), TOPP6 (At5g43380), TOPP7 (At4g11240),
TOPP8 (At5g27840), TOPP9 (not generally referred to as TOPP9, but listed as this here for clarity; At3g05580), AtPP2A-1 (At1g59830), AtPP2A-2 (At1g10430), AtPP2A-3 (At3g58500), AtPP2A-4
(At2g42500), AtPP2A-5 (At1g69960), AtPP4-1 (At4g26720), AtPP4-2 (At5g55260), AtPP5 (At2g42810), AtPP6 (At1g50370), AtFYPP3 (At3g19980) and AtPP7 (At5g63870). The human sequences
are (GenBank® accession numbers): HsPP1α (NP_002699), HsPP1β (NP_002700), HsPP1γ (NP_002701), HsPP2A-α (NP_002706), HsPP2A-β (NP_004147), HsPP2B-α (NP_000935),
HsPP2B-β (NP_066955) and HsPP2B-γ (NP_005596), HsPP4 (NP_002711), HsPP5 (NP_006238), HsPP6 (NP_002712), HsPP7-1 (PPEF1; NP_006231) and HsPP7-2 (PPEF2; NP_006230).
eukaryota is probably due to its role in the dephosphorylation of
the CTD of RNA polymerase II, a process that is itself ancient
and conserved. This is in contrast with the EYA (eyes absent)
c The Authors Journal compilation c 2009 Biochemical Society
protein, another aspartate-based phosphatase, which has
been identified only in multicellular eukaryotes (one to four
members per organism), and plays a role in development.
Protein phosphatase evolution
Mutations of these proteins lead to severe developmental
abnormalities, such as Branchio-oto-renal syndrome, caused
by mutation to the human EYA1 protein [79]. Most EYA
homologues contain an N-terminal transactivation domain that
forms an active transcription factor along with orthologues of
the SIX (Sine oculis) protein of Drosophila. The exception to
this rule is in plants, where Arabidopsis and Oryza express EYA
orthologues that lack transactivation domains [27]. The absence
of the transactivation domain itself is not surprising, as plants lack
a homologue to SIX. However, since plants also lack all of the
systems currently known to be affected by mutation of the EYA
proteins, the function of these proteins in plants is still unknown.
The aspartate-based phosphatases provide two excellent
examples of how protein sequence conservation can infer functional conservation. The first is a ubiquitin-like domain containing
CTD phosphatase, UBLCP1, which, like the EYA proteins, is
conserved throughout multicellular eukaryotes. Only a single
study exists on the human protein, demonstrating both phosphatase activity and nuclear localization of UBLCP1 [80].
The second is a 50 kDa translocase of the inner mitochondrial
membrane (termed TIM50 or TIMM50), an essential component
of the mitochondrial inner membrane in both human and
yeast cells [81]. This protein has been studied in some detail;
however, very little attention has been given to the presence
of the phosphatase domain, save a single study demonstrating
protein phosphatase activity [81]. Simply due to their respective
conservation patterns, we infer that UBLCP1 will be involved, as
are the EYA proteins, with multicellular development, and that
the phosphatase domain of TIM50 plays an important role in the
regulation of mitochondrial translocation.
Conversely, where species have ‘unique’ versions of these
proteins, it suggests a species-specific role. Arabidopsis, rice and
poplar genomes indicate a dramatic expansion in the total number
of FCP-like phosphatases, with more than double the number of
proteins compared with humans (19, 19 and 23 respectively,
compared with eight in humans [27]). Hence these proteins may
have a number of unique roles in plants, indeed demonstrated by
recent studies implicating two of the unique members, CPL1 and
CPL2, in jasmonic acid biosynthesis, as well as stress and auxin
responses [82,83].
CONVERGENCE ON FUNCTION: EVOLUTION OF
CARBOHYDRATE-BINDING PHOSPHATASES
Two of the best examples of convergent evolution at the molecular level come from the phosphatase field. We have already
noted the structural convergence of the PPP and PPM active
sites with little or no sequence relatedness in their respective
genes. More recently the human and plant PTPs, laforin and
SEX4 (starch excess 4), have been demonstrated to be functional
equivalents [84]. Lafora disease is an autosomal recessive
disorder characterized by the accumulation of long-stranded,
poorly branched glycogen (polyglucosan), known as Lafora
bodies. Nearly 50 % of Lafora disease patients have a recessive
mutation in EPM2A (epilepsy of progressive myoclonus type 2
gene A) [85] that encodes laforin, a DSP with a carbohydratebinding module (CBM 20) N-terminal to the PTP domain
(Figure 3). Laforin is the only human phosphatase with a
carbohydrate-docking module (PP1 associates with glycogen
through separate glycogen-binding subunits) and displays the
currently unique ability among PTPs to dephosphorylate complex
carbohydrates [84,86]. Consistent with results from a mouse
knockout of EPM2A, laforin is now recognized as a glycogen
phosphatase and mutation of the laforin gene leads to aberrant
Figure 3
407
Domain structure and gene organization of SEX4 and laforin
The plant starch phosphatase SEX4 is shown with its chloroplast transit peptide (cTP),
dual-specificity (or PTP) phosphatase domain (DSP) and carbohydrate-binding module 20
(CBM20). Human laforin has dual-specificity (or PTP) phosphatase domain (DSP) and
carbohydrate-binding module 20 (CBM20) domains, but they are reversed in order indicating
convergent evolution to a common target substrate.
accumulation of glycogen and disease development. Both starch
and glycogen are phosphorylated molecules and only recently
has the significance of this modification been appreciated. In
plants the phosphorylation and dephosphorylation of starch is critical to its accumulation and appropriate mobilization in leaves
[87–89]. The power of genomics combined with phylogenetics
(and biochemistry) was nicely demonstrated when laforin was
further investigated [84]. A number of protists (unicellular eukaryotes) synthesize and store floridean starch, another complex
carbohydrate related to amylopectin. Genomics of several of
these protists (Toxoplasma gondi, Eimeria tenella, Tetrahymena
thermophila, Paramecium tetraurelia and Cyanidioschyzon merolae) has revealed putative laforin orthologues. Characterization
of the least-conserved laforin (from C. merolae) showed it has
biochemical properties similar to human laforin and localizes to
floridean starch in vivo [84]. Each of these five protists identified
with a laforin orthologue are of red algal descent, contain a true
mitochondrion and produce floridean starch. Other organisms of
red algal descent lack laforin orthologues [88] and in each case
they were noted to not make floridean starch and/or not have
true mitochondria. Further analysis of genomic data revealed
that laforin orthologues only exist in these five protists and all
vertebrates, but not in any invertebrate animals or other eukaryote,
bacteria or archeal genomes. In the same study it was suggested
that laforin originated in an ancestral red algae early in eukaryotic
evolution and was maintained only in vertebrates and those
organisms that produce floridean starch.
A genetic screen for plants that produce excess starch (SEX
mutants) yielded several novel mutants including one designated
SEX4 whose locus was mapped to At3g52180 in Arabidopsis.
This gene, like laforin, encodes a DSP with a carbohydratebinding module (CBM 20), but in this case the domain order
is reversed (Figure 3) indicating that SEX4 and laforin are
not orthologues. SEX4 was predicted and shown to localize to
the chloroplast (the site of starch synthesis/degradation) and to associate with and dephosphorylate starch [87–89]. Being the only
genes in animals and plants whose mutation causes inappropriate
glycogen/starch accumulation in combination with both having
CBM and DSP domains, led Gentry et al. [84] to postulate that
SEX4 and laforin were functional equivalents. In an elegant study
they generated stable cell lines of the Arabidopsis SEX4 mutant
cell line sex4-3 transformed with either wild-type SEX4 or human
laforin (preceded by a SEX4 chloroplast transit peptide). Both of
these reverted the sex4-3 mutant starch-excess phenotype. This is
an excellent example of convergent evolution showing that plants
and animals evolved functionally equivalent phosphatases.
CONCLUSION
The age of comparative genomics will have a major impact on
evolutionary biology and biological research in general. Already
c The Authors Journal compilation c 2009 Biochemical Society
408
G. B. G. Moorhead and others
this field has played a role in resolving key issues in the
phylogenetic relationships between organisms (particularly those
that sit at evolutionary nodes) and shedding light on the emergence
of basic cellular processes [90] and evolution of species-specific
traits [91].
Information gleaned from genomics studies will also allow new
large-scale or systems biology approaches to answer research
questions in all fields of biology. For example, knockdown and
knockout studies can be initiated that take a complement of
phosphatases and asks what phenotype is generated when the
expression of an individual enzyme is dramatically modified or
eliminated. We can also do experiments where individual enzymes
from the phosphatase catalogue can be overexpressed in cells
and then examine their influence on the phosphorylation state of
particular proteins. Indeed, the age of comparative genomics has
meant that the protein phosphatases have come of age.
FUNDING
This work was supported by the Natural Sciences and Engineering Research Council of
Canada [grant number 216895]; the Alberta Ingenuity Center for Carbohydrate Science
[grant number RT707047]; and the Alberta Cancer Board [grant number 23161].
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Received 3 October 2008/18 November; accepted 19 November 2008
Published on the Internet 23 December 2008, doi:10.1042/BJ20081986
c The Authors Journal compilation c 2009 Biochemical Society
Biochem. J. (2009) 417, 401–409 (Printed in Great Britain)
doi:10.1042/BJ20081986
SUPPLEMENTARY ONLINE DATA
Evolution of protein phosphatases in plants and animals
Greg B. G. MOORHEAD1 , Veerle DE WEVER, George TEMPLETON and David KERK
Department of Biological Sciences, University of Calgary, 2500 University Dr., N.W. Calgary, Alberta, Canada T2N 1N4
Figure S1
Phylogenetic analysis of the PPP family kelch-domain containing phosphatases
A rooted tree was generated by comparison of PPKL sequences derived from tBLASTn searches throughout eukaryotic genomes. Sequences were aligned using ClustalX, hand-optimized using
GeneDoc, and a bootstrap tree was generated with the neighbour-joining function of ClustalX [1]. Sequences are derived from Uniprot/TrEMBL (http://www.expasy.org) unless stated otherwise:
Ostreococcus lucimarinus , A4S3K6; Chlamydomonas reinhardtii , A8HNE3; Physcomitrella patens a , A9S2C2; Physcomitrella patens b , A9STN4; Physcomitrella patens c , A9SF24; Physcomitrella
patens d , A9TQY8; Arabidopsis thaliana , At1g03445, Q9LR78; At1g08420, Q9SJF0; At2g27210, Q9SHS7; At4g03080, Q9ZTA4; Vinis Vitifera a , A7QVR2; Vinis Vitifera b , A7PU12; Oryza sativa
indica a , A2XK43; Oryza sativa indica b , A2ZMT0; Oryza sativa japonica a , A3AKU8; Oryza sativa japonica b , A3BCU1; Oryza sativa japonica c , Q2QM47; Oryza sativa japonica d , Q10G19;
Oryza sativa japonica e , Q60EX6; Solanum lycopersicum (NCBI nucleotide database: BT013516); Babesia bovis , A7AT59; Plasmodium falciparium , Q8IKH5 identical with O96914; Plasmodium
berghei , Q4Z2M2; Plasmodium knowlesi , B3L999; Plasmodium chabaudi , Q4XN17; Plasmodium vivax ; A5K396; Plasmodium yoelii yoelii , Q7RA97; Theileria parva , Q4N2C8; Theileria annulata ,
Q4U9P3; Tetrahymena thermophila a (NCBI protein XP_001031447); Tetrahymena thermophila b , Q240X8; Tetrahymena thermophila c , Q22BC4; Neospora caninum (GeneDB NC_LIV_113060);
Cryptosporidium hominis , Q5CHL7; Cryptosporidium parvum , A3FQS0; Paramecium tetraurelia a , A0BV09; Paramecium tetraurelia b , A0BZY8; Paramecium tetraurelia c , A0CLB3; Paramecium
tetraurelia d , A0CV78; Paramecium tetraurelia e , A0DWN0; Paramecium tetraurelia f , A0D9U9; and Paramecium tetraurelia g , A0E8D8. The human PP1α sequence P62136 was used as an outgroup.
1
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2009 Biochemical Society
G. B. G. Moorhead and others
Figure S2
For legend see last page.
c The Authors Journal compilation c 2009 Biochemical Society
Protein phosphatase evolution
Figure S2
For legend see last page.
c The Authors Journal compilation c 2009 Biochemical Society
G. B. G. Moorhead and others
Figure S2
For legend see last page.
c The Authors Journal compilation c 2009 Biochemical Society
Protein phosphatase evolution
Figure S2
For legend see last page.
c The Authors Journal compilation c 2009 Biochemical Society
G. B. G. Moorhead and others
Figure S2
Multiple sequence alignment of protein phosphatases with kelch domains
For sequence annotations and alignment method, see the legend of Figure 2 in the main text. Kelch domains are indicated above according to [2]. Conserved residues within are marked with $.
Consensus secondary structure predictions obtained from (http://npsa-pbil.ibcp.fr/) with default parameters were indicated: α-helices are shown in bold and β-strands are underlined. Indecisive
predictions are bold and underlined. Inserts in the PPKL phosphatase domains, as compared with the PPP catalytic domains, are indicated above according to [2]. Conserved residues within
the phosphatase domain, compared with human PP1α (!) or PPP family members (*) [3] are indicated with symbols shown between parentheses while residues involved in inhibitor (#) ([4] and
V. DeWever and G. Moorhead, unpublished work) and RVXF-motif binding (%) [4,5] defined based on PP1 studies are marked as indicated between parentheses.
c The Authors Journal compilation c 2009 Biochemical Society
Protein phosphatase evolution
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phosphatases. Eur. J. Biochem. 220, 225–237
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Nuclear protein phosphatases with Kelch-repeat domains modulate the response to
brassinosteroids in Arabidopsis . Genes Dev. 18, 448–460
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targeting subunit binding. Biochemistry 40, 7410–7420
Received 3 October 2008/18 November; accepted 19 November 2008
Published on the Internet 23 December 2008, doi:10.1042/BJ20081986
c The Authors Journal compilation c 2009 Biochemical Society