Biochem. J. (2008) 409, 333–348 (Printed in Great Britain)
333
doi:10.1042/BJ20071097
REVIEW ARTICLE
The histidine phosphatase superfamily: structure and function
Daniel J. RIGDEN1
School of Biological Sciences, University of Liverpool, Crown Street, Liverpool L69 7ZB, U.K.
The histidine phosphatase superfamily is a large functionally
diverse group of proteins. They share a conserved catalytic core
centred on a histidine which becomes phosphorylated during the
course of the reaction. Although the superfamily is overwhelmingly composed of phosphatases, the earliest known and arguably
best-studied member is dPGM (cofactor-dependent phosphoglycerate mutase). The superfamily contains two branches sharing
very limited sequence similarity: the first containing dPGM,
fructose-2,6-bisphosphatase, PhoE, SixA, TIGAR [TP53 (tumour
protein 53)-induced glycolysis and apoptosis regulator], Sts-1 and
many other activities, and the second, smaller, branch composed
mainly of acid phosphatases and phytases. Human representatives
of both branches are of considerable medical interest, and
various parasites contain superfamily members whose inhibition
might have therapeutic value. Additionally, several phosphatases,
notably the phytases, have current or potential applications in
agriculture. The present review aims to draw together what is
known about structure and function in the superfamily. With the
benefit of an expanding set of histidine phosphatase superfamily
structures, a clearer picture of the conserved elements is obtained,
along with, conversely, a view of the sometimes surprising
variation in substrate-binding and proton donor residues across
the superfamily. This analysis should contribute to correcting a
history of over- and mis-annotation in the superfamily, but also
suggests that structural knowledge, from models or experimental
structures, in conjunction with experimental assays, will prove
vital for the future description of function in the superfamily.
INTRODUCTION
in the family is essential to inform future function annotation and
is one of the objectives of the present review. Another is to bring
together, for the first time, the literature on activities of histidine
phosphatases and thereby illustrate how impressively diverse they
are, involving many proteins of medical or applied interest.
Research into the histidine phosphatase superfamily dates back to
1935 when an activity was found in yeast capable of interconverting 2PGA and 3PGA (2- and 3-phosphoglycerate) [1]. Of course,
it was only later that the sequence of the enzyme responsible,
dPGM (cofactor-dependent phosphoglycerate mutase), was determined [2], and later still when it was realized that very different
activities, the first being F26BPase (fructose-2,6-bisphosphatase),
could be catalysed by homologous enzymes [3]. In the 20 years
since, a great many different functions have been associated with
the superfamily and structures have been determined for some
of the respective proteins. All of these new discoveries have been
phosphatase activities of one kind or another, illustrating that,
ironically enough, the earliest and arguably best studied member
of the superfamily is anomalous in catalysing primarily a mutase
reaction. Although the superfamily has undoubtedly benefited
from containing the key glycolytic enzyme dPGM, the subject of
many early studies [4], a reading of the literature also illustrates
a disadvantage arising from the mutase-dominated superfamily
history; many superfamily members discovered in genomes have
been suggested to be mutases when a phosphatase activity
is in fact much more likely. Indeed, like other superfamilies
[5], the histidine phosphatases present considerable challenges
to automated genome annotation tools and have a history of
widespread mis- and over-annotation. Even human interpretation
of sequence relationships has been regularly biased towards
mutase activity rather than the actual predominant phosphatase
activity [6–8]. An analysis of the structure–function relationship
Key words: acid phosphatase, cofactor-dependent phosphoglycerate mutase, fructose-2,6-bisphosphatase, histidine phosphatase,
structure–function relationship.
SEQUENCES
Iterative database searches in current databases using well-known
characterized proteins such as dPGMs or F26BPases provide
clear evidence for a deep split of the superfamily into two
branches. PSI-BLAST searches using queries from branch 1
fail to produced branch 2 members among hit lists and vice
versa. As discussed in detail below, the functions associated
with branch 1 are much more diverse than those in branch 2,
the latter containing mainly enzymes characterized as APs
(acid phosphatases) or phytases (Table 1). Nevertheless, modern
profile–profile comparison methods readily detect the distant
evolutionary relationship between the two branches, a relationship
revealed unambiguously by the first crystal structure of a branch 2
enzyme in 1993 [9]. In fact, even early sequence studies predicted
a relationship between dPGMs and F26BPases (in branch 1) and
APs (in branch 2) on the basis of their sharing a small RHG motif,
near the beginning of the sequence, whose histidine residue was
already known to be phosphorylated during catalysis [10].
Sequences in the histidine phosphatase superfamily are very
diverse, both within and between the two principal branches.
Supplementary Table S1 (http://www.BiochemJ.org/bj/409/
bj4090333add.htm) shows percentage sequence identities
Abbreviations used: AP, acid phosphatase; 1,3-BPG, 1,3-bisphosphoglycerate; 2,3-BPG, 2,3-bisphosphoglycerate; BPGM, bisphosphoglycerate
mutase; dPGM, cofactor-dependent phosphoglycerate mutase; EPP, ecdysteroid phosphate phosphatase; ER, endoplasmic reticulum; F26BP, fructose 2,6bisphosphate; F26BPase, fructose-2,6-bisphosphatase; HPt, histidine-containing phosphotransfer; Minpp1, multiple inositol-polyphosphate phosphatase
1; PAP, prostatic acid phosphatase; PFK-2, phosphofructokinase 2; 2PGA, 2-phosphoglycerate; 3PGA, 3-phosphoglycerate; PGAM5, phosphoglycerate
mutase 5; SH3, Src homology 3; TFIIC, transcription factor IIC; TIGAR, TP53 (tumour protein 53)-induced glycolysis and apoptosis regulator; TRAP,
tartrate-resistant acid phosphatase; UBASH, ubiquitin-associated and SH3 domain-containing.
1
email [email protected]
c The Authors Journal compilation c 2008 Biochemical Society
334
Table 1
D. J. Rigden
The principal known or reliably inferred molecular functions of histidine phosphatase domains grouped into broad functional categories
Detail on reactions catalysed, their broader significance and further references can be found in Supplementary Table S2 (http://www.BiochemJ.org/bj/409/bj4090333add.htm) and in the text.
Branch
1
Catalytic
Broad functional category
Protein and key molecular function reference
Example species
EC number
Intermediary metabolism
dPGM [4]
BPGM [120]
α-Ribazole phosphatase (CobC) [16]
Mannitol-1-phosphatase [79]
Neo13 [18]
PFK2/F26BPase [121]
TIGAR [28]
SixA [15]
E. coli , S. cerevisiae , human
Human
Salmonella Typhimurium
Eimeria tenella
Streptomyces fradiae
Eukaryotes
Eukaryotes
E. coli
5.4.2.1 (also 3.1.3.13 and 5.4.2.4)
5.4.2.4 (also 5.4.2.1 and 3.1.3.13)
3.1.3.73
3.1.3.22
3.1.3.3.1.3.46
3.1.3.46
3.1.3.-
EPP [19]
3.1.3.-
Sts-1, Sts-2 (also known as UBASH3A, TULA) [20]
PMU1 [8]
PhoE [116]
TFIIIC τ 55 kDa subunit [39]
Bombyx mori , Drosophila
melanogaster
Human
S. cerevisiae
G. stearothermophilus
S. cerevisiae
PGAM5 [38]
Pho5p [22]
Human
S. cerevisiae
May include 3.6.1.3
Glucose-1-phosphatase [21]
Phytase [23]
Minpp1 [122]
E. coli
Aspergilli and E. coli
Human
3.1.3.10
3.1.3.8, 3.1.3.26
3.1.3.62
Lysophosphatidic acid phosphatase [30]
PAP [123]
Human
Human
3.1.3.2
3.1.3.48
Testicular acid phosphatase [34]
Lysosomal acid phosphatase (LAP)
and Acp5/TRAP [36]
Peripheral vacuolar acid phosphatase [37]
Human
Human
3.1.3.48
?
Giardia lamblia
?
Metabolic regulation
Other regulatory or
developmental process
Unknown
Non-catalytic
(?)
2
Catalytic
Extracellular
metabolism/scavenging
Regulation of signalling
molecule concentration
Other regulatory or
developmental process
determined for pairwise comparisons between members of known
structure. In branch 1, SixA, which is considerably smaller
than the other members (see below), shares only 15–19 %
sequence identity with them. The similarly sized dPGM, PhoE and
F26BPase share just 19–28 % sequence identity between different
catalytic activities. The comparison of dPGMs and BPGMs
(bisphosphoglycerate mutases) shows how simple sequenceidentity-based function annotation can easily go astray: human
dPGM and BPGM, with different principal catalytic activities,
share 53 % sequence identity, whereas the value for the bacterial
dPGMs from Escherichia coli and Mycobacterium tuberculosis,
catalysing the same reaction, is only 42 %. Between branches 1
and 2 the highest observed percentage sequence identity is just
18 %. Low values, down to 9 %, are also recorded for pairwise
comparisons within branch 2 where less well conserved N- and
C-terminal extensions can contribute to diversity.
The division of the superfamily is reflected in its twin entries in
the Pfam domain database [11] (http://pfam.sanger.ac.uk/). The
‘phosphoglycerate mutase family’ entry (PF00300) corresponds
to branch 1 and the ‘histidine acid phosphatase’ entry (PF00328)
represents branch 2. Their relationship is acknowledged by their
union in the ‘phosphoglycerate mutase-like superfamily’ clan
(CL0071). In the present version of the Pfam database (version
22, July 2007), branch 1 with just over 3000 members outnumbers
branch 2 by four to one. Interestingly, archaea contain no members
of branch 2 and just 37 representatives of branch 1. These
numbers, and the diversity of the archaeal sequences, suggest that
the common ancestor of archaea lacked a histidine phosphatase
and a small number of organisms later acquired branch 1
members by lateral gene transfer. Interestingly, branch 2 proteins
are largely found in eukaryotes (634 sequences) with fewer in
c The Authors Journal compilation c 2008 Biochemical Society
3.1.3.3.1.3.3.1.3.-
bacteria (108 sequences). In branch 1, the reverse is true, with
bacterial sequences (2192) outnumbering eukaryotic proteins
(781).
The COG (Clusters of Orthologous Groups) database [12]
(http://www.ncbi.nlm.nih.gov/COG/), designed to group orthologous proteins from genome sequences, covers a small portion of
the histidine phosphatase superfamily with three entries. However,
there are a number of problems with these entries, problems that
automated annotation pipelines have unfortunately propagated
into large numbers of individual sequence annotations. For
example, examination of COG0588, ‘phosphoglycerate mutase
1’, shows that many members do not share substrate-binding
residues (discussed later) with the well-characterized E. coli
dPGM and therefore are probably not genuine dPGM enzymes.
Worse, COG0406, containing predominantly bacterial sequences,
is labelled ‘fructose-2,6-bisphosphatase’ on the basis of the
inclusion of a Saccharomyces cerevisiae sequence with that
activity, yet F26BP has not been detected outside eukaryotes. {A
putatively active PFK-2 (phosphofructokinase 2)/F26BPase has
been found in two Desulfovibrio species [13,14], neither present in
COG0406.} The remaining entry is COG2062, ‘phosphohistidine
phosphatase SixA’.
Importantly, many members of the superfamily cannot be
reliably annotated with any of the known functions (see below)
since they bear very limited sequence similarity to characterized
members. Clustering experiments (D. J. Rigden, unpublished
work) show many well-defined groups, presumably sharing the
same function, for which no functional data are available. Given
sufficient knowledge of the structure–function relationship in the
superfamily, it should be possible to apply bioinformatics tools to
predict their functions.
Structure and function in the histidine phosphatase superfamily
FUNCTIONS
A census
Supplementary Table S2 (http://www.BiochemJ.org/bj/409/
bj4090333add.htm) presents a compilation of functions
characterized for members of the histidine phosphatase
superfamily. It contains only activities, demonstrated or strongly
inferred, with potential physiological relevance: many enzymes
will hydrolyse unnatural model substrates, for example. A
condensed version with activities placed in broad categories is
shown as Table 1. The most impressive feature of the superfamily
is the range of activities catalysed. This is manifest most obviously
in the presence of the dPGMs in a list otherwise dominated by
phosphatases. As discussed later, a common catalytic machinery
is shared by mutases and phosphatases. The list also shows
that specificities among enzymes vary dramatically from small
molecules such as phosphoglycerate (M r = 187), the substrate
of dPGM, to large phosphorylated proteins such as the HPt
(histidine-containing phosphotransfer) domain of ArcB [15], the
substrate of SixA (M r = 9050).
dPGM, as a participant in both glycolysis and gluconeogenesis,
contributes to both catabolic and anabolic pathways. Other
anabolic enzymes include CobC, involved in cobalamin synthesis
[16], mannitol-1-phosphatase, which forms the Eimeria parasite
storage compound mannitol [17], and Neo13, characterized
from Streptomyces fradiae, which contributes to synthesis
of the antibiotic neomycin [18]. Catabolic enzymes include
EPP (ecdysteroid phosphate phosphatase), which releases free
ecdysteroids from conjugated reserves in insects [19,20], and
glucose-1-phosphatase, required for growth of E. coli on a glucose
1-phosphate carbon source [21]. Fungi have two superfamily
members involved in scavenging phosphate from extracellular
sources: Pho5p, which degrades extracellular nucleotides [22],
and phytase, which releases the phosphate groups from phytic
acid [23,24].
As well as metabolic functions, a surprisingly large proportion
of the family is involved in signalling or regulation of one kind
or another. EPP, mentioned above, is likely to be important
for insect development, whereas two distinct members of the
family, F26BPase and TIGAR [TP53 (tumour protein 53)-induced
glycolysis and apoptosis regulator], have roles, demonstrated or
strongly suggested respectively, in controlling the concentration
of intracellular F26BP (fructose 2,6-bisphosphate) and thereby
influencing the rates of glycolysis and gluconeogenesis in
eukaryotic cells [25–27]. For a fuller discussion of F26BPases,
and their N-terminal PFK-2 domains, the reader is referred
to other reviews [13,14]. Interestingly, the well-characterized
F26BP-binding residues of F26BPase are not conserved in
TIGAR, indicating that the two proteins have converged on
the same substrate and share more distant ancestry than their
common activity would suggest [28]. The primary in vivo role of
PMU1 (whose name derives from mis-annotation as a potential
phosphomutase) in yeast is likely to involve regulation of the
concentration of another small molecule, AICAR (5-amino-4imidazolecarboxamide riboside), and which has an influence over
transcription of adenine synthesis genes [8]. Eukaryotic Minpp1
(multiple inositol-polyphosphate phosphatase 1) regulates the
cellular concentrations of other signalling molecules, namely
inositol pentakisphosphate and inositol hexakisphosphate [29].
Also in eukaryotes, the signalling molecule lysophosphatidic acid
is a known substrate of both lysophosphatidic acid phosphatase
[30] and PAP (prostatic acid phosphatase) [31].
Protein phosphatases from both branches have regulatory roles.
In branch 1, SixA acts on a phosphohistidine residue in ArcB
with a well-characterized role as a regulator of two-component
335
signalling in bacteria [15,32]. In branch 2, superfamily members
target phosphotyrosine residues in important regulatory proteins
such as ErbB2 [33], ErbB4 [34] and the EGF (epidermal growth
factor) receptor [35]. Other protein phosphatase activities in
branch 2 have interestingly different roles, in vertebrate bone
development [36] and in excystation of the parasite Giardia
lamblia [37].
In two cases, eukaryotic members of the superfamily have
had wholly non-catalytic roles proposed for them: protein–
protein interactions in the case of human PGAM5 [38] and
stabilization/assembly in the case of the TFIIIC (transcription
factor IIIC) τ 55 kDa subunit in yeast [39]. However, in both
cases, the prerequisites for catalysis (see below) are apparently
present so that catalytic roles remain possible. This situation is
graphically illustrated by the story of Sts-1, a key T-cell regulator
[6,40,41] with a multidomain structure. The presence of a histidine
phosphatase domain led investigators to test for dPGM activity
and find none [6]. Since loss of that domain destroyed the dimeric
quaternary structure, it was proposed that it functioned merely to
assemble the dimer [6]. However, after its close kinship with EPP
was recognized, it was tested for and was found to have activity
against ecdysteroid and steroid phosphates [20], although its true
in vivo substrate may be neither.
There are several cases where indirect evidence provides broad
clues as to the function of histidine phosphatase superfamily members. For example, the cell-surface Pho4 protein of Schizosaccharomyces pombe can be repressed by thiamine, suggesting a
role in thiamine metabolism, possibly cleavage of a thiamine
phosphate [42]. In the case of BluF from Rhodobacter capsulatus,
it is the genomic location of its gene in an operon containing
exclusively cobalamin synthesis genes which argues strongly for
a role (distinct from CobC above, also present in R. capsulatus) in
that process [43]. Similarly, ORF2 (open reading frame 2) of the
plasmid-borne avrPphF gene in Pseudomonas syringae pathovar
phaseolicola, a histidine phosphatase, was implicated in bacterial
virulence, a role later confirmed, although its substrate remains
unknown [44]. For a homologue from Arabidopsis thaliana, At74,
induction by glucose and an increase in glucose uptake by overexpressing plants point to a role in carbohydrate metabolism [45].
Domain fusions are among the most potent sources of
functional clues in the superfamily. Figure 1 illustrates the domain
content of at least partly characterized members of the histidine
phosphatase superfamily. Genome searches suggest that further
functionally suggestive domain architectures exist in other parts
of the superfamily (D. J. Rigden, unpublished work). First among
these domain combinations is the pairing of F26BPase with the
kinase domain (placed N-terminally) that catalyses the nearreverse reaction: synthesis of F26BP. The relative activity of these
two domains is regulated by phosphorylation state in a way not
yet fully understood [14], and differs between mammalian isoenzymes due to intrinsic sequence features [46]. Very recently, a potentially similar situation has been discovered involving a branch
2 histidine phosphatase domain [47]. The N-terminal domain of
yeast Vip1 has inositol-hexakisphosphate kinase activity and is
followed by a phosphatase domain of as yet unknown specificity,
but which is very likely to act on multiply phosphorylated inositol
species. The SH3 (Src homology 3) and UBASH (ubiquitinassociated and SH3 domain-containing) domains found in EPP
and Sts-1, suggestive of protein- and ubiquitin-binding roles respectively, have been investigated experimentally [6], confirming
predictions. The same proteins also contain 2H phosphoesterase
domains [48], as yet uncharacterized, but likely to share ligands
in some way with the histidine phosphatase domain. Additional
domains have been noted in some PFK-2/F26BPase sequences
[13,49]. Thus, in some trypanosomatid sequences, multiple
c The Authors Journal compilation c 2008 Biochemical Society
336
Figure 1
D. J. Rigden
Domain architectures of selected histidine phosphatase superfamily members
All domain combinations that are at least partially characterized are shown, as well as some designed to illustrate the range of sizes of phosphatase domains. Domains are drawn approximately to
scale. Catalytic domains are coloured blue (histidine phosphatase domains; dark for branch 1, light for branch 2), red (PFK-2 domains), purple (the 2H phosphoesterase domain seen in Sts-1 and
relatives [20]), pink (the reductase domain of enzymes involved in opine synthesis [50–52]) and brown (the Vip1 kinase domain [47]). Other domains are coloured mauve (the carbohydrate-binding
module of plant PFK-2/F26BPases [13]), green (the ankyrin repeats seen in some trypanosomatid PFK-2/F26BPases [49]; darker shades indicating more reliably assigned repeats), and grey [UBA
(ubiquitin-associated)] or yellow (SH3), both found in Sts-1 and relatives [20].
ankyrin repeat motifs are present [49], indicating a possible
site of interaction of the PFK-2/F26BPase protein with others,
while plant homologues contain a carbohydrate-binding module
at their N-terminus, suggesting a possible means to control PFK-2/
F26BPase activity [13]. Finally, histidine phosphatase domains
are found N-terminally fused to dehydrogenase domains in
proteins involved in opine synthesis [50–52]. With the C-terminal
domains assigned functions such as agropine synthesis reductase
[50], it is clear that the histidine phosphatase domains will be
involved in some way in the opine synthetic pathway.
Subcellular localization
When the known subcellular localizations of histidine
phosphatase superfamily members are listed, an interesting
dichotomy is observed. In branch 1, cytoplasmic location is the
norm with other members found partially or wholly in the nucleus
(Table 2). The only known exceptions to this rule are the predicted
periplasmic location of the Ais/AfrS family [53], and the presence
in yeast cell wall of dPGM [54]. In contrast, all branch 2 family
members appear to enter the secretory pathway. Some remain
in the ER (endoplasmic reticulum), others are found at the cell
surface, periplasm or cell wall, and others are simply secreted
(Table 2). Lysosomal AP, unusually, is transported as a type I
membrane protein from the ER, via the plasma membrane, to the
c The Authors Journal compilation c 2008 Biochemical Society
lysosome [55] where its luminal portion is slowly proteolytically
cleaved.
There seems to be no obvious intrinsic reason for the largely
distinct subcellular localizations of the two branches, but the fact
that the division is visible across bacteria and eukaryotes perhaps
suggests that each branch became specialized for an intra- or
extra-cellular use soon after an early divergence. In accordance
with their subcellular localization, branch 2 enzymes commonly,
although not invariably, contain disulfide bonds, with Swiss-Prot
[56] entries cataloguing anything from zero to five of them per
protein. This range is confirmed by available crystal structures,
since an enzyme from Francisella tularensis (PDB code 2GLC)
has none, whereas Aspergillus fumigatus phytase (PDB code
1QWO) has five. The disulfide bonds, where present, are found
throughout the structure, but not near the catalytic site, indicating
further a general stabilizing function rather than any regulatory
role. Swiss-Prot entries indicate no known disulfide bonds in
branch 1, but conservation and proximity in a molecular model
suggest strongly that the unusual periplasmic branch 1 enzymes
of the Ais/AfrS family contain two [53].
Medical and applied importance
As expected from their diverse important roles, deficiencies or
mutations in many human members of the histidine phosphatase
superfamily, from both branches 1 and 2, lead to diseases.
Structure and function in the histidine phosphatase superfamily
Table 2
337
Subcellular localization of some histidine phosphatase superfamily members
Branch
Localization
Protein
Representative species
Evidence for localization
1
Cytosol
dPGM
BPGM
F26BPase
TFIIIC τ 55 kDa subunit
UBASH3A/TULA/SRS-2
PGAM5
dPGM
YOR283W
YDR051C
dPGM
Ais
Human
Rat
Rat
S. cerevisiae
Human
Human
Rat
S. cerevisiae
S. cerevisiae
S. cerevisiae
E. coli
[4]
[124]
[125]
[39]
[6,126]
[38]
[127,128]
[129]
[129]
[54]
Predicted in [53]
Lysophosphatidic acid phosphatase
Multiple inositol-polyphosphate phosphatase
AP
AP
Testicular acid phosphatase
Thiamine-repressible acid phosphatase precursor
Phytase
Glucose-1-phosphatase
Pho5p
Phytase
Human
Human
Leishmania mexicana
Leishmania donovani
Human
S. pombe
E. coli
E. coli
S. cerevisiae
Aspergillus ficuum
[130]
[29,122]
[131]
[132]
Swiss-Prot [56]
[133]
[134]
[21]
[135]
[136]
Nuclear
Cytosol and nuclear
Cytosol and cell wall
Periplasm
2
Lysosome
ER/plasma membrane
Endosomal/lysosomal compartment
Cell membrane (type I, catalytic domain outside cell)
Cell membrane (type I, catalytic domain outside cell)
Cell wall
Periplasm
Secreted
Deficiency conditions of varying degrees of severity result from
loss of activity of dPGM [57], BPGM [58], lysosomal AP [59] or
macrophage AP [60].
For some branch 1 human enzymes, there is potential
therapeutic interest in inhibition of activity. One exciting
emerging medical angle concerns dPGM as a sensitive node of
the glycolytic pathway, which is up-regulated in cancer cells
of various kinds [61]. For example, when phosphorylation of
dPGM was prevented in a tumour cell line, a reduction in
glycolysis and arrest of tumour growth was observed [62]. Later,
a compound MJE3 isolated as having an anti-proliferative effect
on breast tumour cells [63] was found to target dPGM by
covalent attachment of a spiroepoxide group to a lysine residue
near the catalytic site [64]. Modulation of erythrocyte BPGM
activity is also of potential medical interest. BPGM is the sole
enzyme responsible for controlling the concentration of 2,3-BPG
(2,3-bisphosphoglycerate) in erythrocytes. Its synthase activity
produces 2,3-BPG, while the phosphatase side reaction degrades
it (Table 1). 2,3-BPG lowers the apparent affinity of haemoglobin
for oxygen by binding selectively to the deoxygenated form. A
rise in 2,3-BPG concentration is consistently seen in conditions of
hypoxia [65], suggesting an adaptive response while lowering 2,3BPG concentrations has an anti-sickling effect [66]. Since BPGM
is responsible for the loss of 2,3-BPG in stored blood, there is also
an applied interest in discovery of an agent that would inhibit its
phosphatase activity [67].
Although it is fair to say that we still lack a complete
understanding of AP activities and in vivo substrates, these branch
2 enzymes have important clinical applications (reviewed in [68]).
In particular, PAP has long been used as a serum marker for
prostate cancer [69], and its overexpression on cancer cells is
being exploited for immunotherapy [70]. More speculatively, both
well-characterized activities of PAP lead to down-regulation of
cancer growth [31,33], suggesting that the enzyme may have
therapeutic value. Conversely, should inhibition of the enzyme be
desired, good inhibitors are already available [71]. TRAP (tartrateresistant acid phosphatase) is a marker for bone resorption and
can be monitored in patients with metabolic bone disorders
[72]. Again in humans, the suggestion that multiple inositol-
polyphosphate phosphatase could contribute to the pathogenesis
of some thyroid carcinomas [73] means that specific inhibitors
could be of therapeutic interest.
In other medically relevant species, histidine phosphatases are
also of interest. For example, a peripheral vacuolar AP is required
for excystation of the parasite Giardia lamblia [37], suggesting
that a selective inhibitor of that enzyme might have value as an
anti-parasitic agent. In another group of parasites, the Leishmania
species, three APs have been described and, although conclusive
evidence is lacking, their conservation among Leishmania species
is interpreted as suggestive of an important role in the growth or
development of the parasite [74]. Finally, a better understanding of
the molecular functions of the histidine phosphatases implicated
in synthesis of different classes of antibiotics [18,50–52,75] may
help efforts to produce novel antibiotics.
The most well-known applied use of a histidine phosphatase is
the addition of phytase to animal feed, first in order to increase the
availability of the phosphate component of phytate, obviating
the need to supply phosphorus supplements, and secondly to
liberate iron and other trace elements from their phytate chelation
[76]. To this end, phytases with improved molecular properties
have been both designed through mutations of known enzymes
[77] and sought by screening (e.g. [78]). Another potential
agricultural application could involve EPP [19,20] as a novel
insecticide target. Although formal validation is still required,
the role of EPP in liberating inactive stores of ecdysteroids
during insect development suggests that its inhibition should have
serious consequences for the insect. Finally, parasitic protozoa
of the genus Eimeria are a significant problem in the poultry
industry. These organisms use mannitol-1-phosphatase during the
synthesis of mannitol, an essential storage compound, suggesting
that inhibitors of this enzyme, absent from the host, could be
useful for parasite control [79].
STRUCTURES
Table 3 presents a compilation of all histidine phosphatase
superfamily structures available at the time of writing. As the
c The Authors Journal compilation c 2008 Biochemical Society
338
D. J. Rigden
c The Authors Journal compilation c 2008 Biochemical Society
Table 3
Available structures of members of the histidine phosphatase superfamily
1 Å = 0.1 nm
Major
branch
Enzyme activity
Source
Year of deposition
in the PDB
PDB code(s)
Resolution (Å)
Reference(s)
Notes
1
dPGM
S. cerevisiae
1975
1PGM*
3.5
[137,138]
1982
1997
3PGM†
4PGM
2.8
2.3
[139]
[81]
The first superfamily crystal structure, published in 1974 [138] following
an earlier low-resolution structure [137]. No sequence available.
The first sequence-traced structure.
Corrects an error in the conformation of the active-site histidine residues
of earlier structures.
S. pombe
E. coli
1998–1999
2000
2000
5PGM, 1BQ3, 1BQ4, 1QHF
1FZT
1E58, 1E59
1.7–2.7
n/a
1.25–1.3
[107,108,140]
[141]
[89,106]
M. tuberculosis
Plasmodium falciparum
Human (B-type)
Human erythrocyte
2003
2004
2005
2004–2005
1RII
1XQ9
1YJX, 1YFK
1T8P, 2A9J, 2HHJ, 2F90, 2H52, 2H4X, 2H4Z
1.7
2.6
2.7–2.8
1.5–2.5
[142]
Unpublished
[112]
[90,109]
G. stearothermophilus
Rat testis
Human liver
E. coli
M. tuberculosis
T. thermophilus
Rat
Human
Aspergillus ficuum
Aspergillus niger
Aspergillus fumigatus
E. coli
E. coli
Francisella tularensis
2001–2002
1996–2000
2001
2003
2005
2003–2006
1993
1998–2002
1997
1999
2003–2004
1999
2003
2006
1EBB, 1H2E, 1H2F
1BIF, 2BIF, 3BIF, 1FBT, 1TIP, 1C80, 1C7Z, 1C81
1K6M
1UJB, 1UJC
2A6P
1V37, 2HIA, 1V7Q
1RPA, 1RPT
2HPA, 1CVI, 1ND5, 1ND6
1IHP
1QFX
1QWO, 1SKA, 1SKB, 1SK9
1DKL, 1DKM, 1DKN, 1DKO, 1DKP, 1DKQ
1NT4
2GLA, 2GLB, 2GLC
1.69–2.3
2.0–2.6
2.4
1.9–2.06
2.2
1.4–1.6
3.0
2.4–3.2
2.5
2.4
1.5–1.69
2.05–2.4
2.4
1.65–1.85
[92,114]
[100,143–146]
[46]
[85]
[86]
Unpublished
[9,82,148]
[71,149,150]
[151]
[152]
[93,115]
[103]
[91]
Unpublished
BPGM
PhoE
F26BPase
2
SixA
Phosphatase of unknown specificity
Unknown
PAP
Phytase
Glucose-1-phosphatase
Unknown
*Obsolete, no longer available in the regular PDB.
†Obsolete, but still present in the PDB.
The only NMR structure of a superfamily member.
Includes the highest resolution structure in the superfamily,
phosphorylated enzyme [89].
Includes a remarkable set of structures providing snapshots along the
process of enzyme phosphorylation by 2,3-BPG [90].
Includes the first structure of a phosphoenzyme in the superfamily [144].
Representative of the minimal core fold of the superfamily.
Unpublished Structural Genomics outputs [147].
The first, chloride-bound structure is not apparently deposited in the PDB.
Includes complexes relevant to rational inhibitor design.
Include a high-resolution structure of the phosphorylated enzyme.
Structure and function in the histidine phosphatase superfamily
Figure 2
339
Consensus tree derived from analysis of all histidine phosphatase superfamily structures
The Figure was generated at the PROCKSI server (http://www.procksi.net). References for the protein structures are shown in Table 3.
deposition dates show, structural studies date back to the early
days of protein crystallography. The first dPGM structure was
deposited in 1975, 1 year after publication of the first Protein Data
Bank [80] newsletter listed just 12 proteins for which co-ordinates
were available. Unfortunately, erroneous positioning of catalytic
histidine residues misled efforts to understand the mechanism [4]
until the availability of a corrected structure in 1997 [81]. The
first branch 2 crystal structure was that of rat PAP in 1993 [82].
In recent years, structure determinations have accelerated, so that
at least nine distinct molecular functions are now covered. In
order to provide a structural context for discussion of catalysis
and substrate diversity, all known structures of the superfamily
were collected together and superimposed with MUSTANG [83]
with post-processing of the results with STACCATO [84]. A
dendrogram representation of the structures, on the basis of a
consensus calculation on the results of six different structural
superposition methods, was created using the PROCKSI server
(http://www.procksi.net) and is shown in Figure 2. This Figure
confirms the broad division of the superfamily into two branches,
the first containing dPGMs, F26BPases and other activities, and
the second containing APs and phytases.
A selection of 11 structures was made in order to represent
the broad diversity of known structures in the superfamily.
Comparison of these, in order of increasing size, with the nearminimal core α/β domain present in SixA [85] reveals how
insertions into the fold differentiate between different lineages and
define the substrate-binding site, thereby conferring the observed
activity. These insertions, sometimes as long as the core domain
itself, together with the sometimes substantial N- and C-terminal
extensions, explain the fact that the largest histidine phosphatase
c The Authors Journal compilation c 2008 Biochemical Society
340
D. J. Rigden
structure yet determined is almost three times the size of the
smallest. PDB accession codes and references can be found in
Table 3. A structure-based sequence alignment of representative
histidine phosphatases is shown in Figure 3.
The principal structural difference between SixA and the
Thermus thermophilus protein of unknown function lies in the 35residue insertion of the latter between β3 and α3. An insertion, of
at least this size, is present here in all structures apart from SixA. In
addition, the β1–α1 loop of the T. thermophilus protein is slightly
larger and adopts a different conformation, packing against the
β3–α3 insertion. The characterized M. tuberculosis phosphatase
[86] resembles the T. thermophilus protein, but has a slightly larger
α4–β5 loop which also packs against the β3–α3 insertion. The
monomeric S. pombe dPGM and Geobacillus stearothermophilus
PhoE (Figure 4b) follow this same general scheme, although,
of course, with detailed sequence differences defining different
specificities (see below). In F26BPase, a substantial 25-residue
C-terminal tail is added to this scheme (Figure 4c). The tail twice
traverses the entrance to the catalytic site, essentially blocking
substrate access, and its removal increases F26BPase activity
[87]. Although a tail is absent from S. pombe dPGM, the E. coli
dPGM structures show how the dPGM C-terminal tails, where
present, adopt a completely different conformation from those in
F26BPases (Figure 4d). dPGM tails are shorter and lie along the
rim of the entrance to the catalytic site. Again with the exception of
the S. pombe enzymes, dPGMs contain an additional ∼ 25-residue
section in the large β3–α3 insertion (Figure 4d), which, in the
tetrameric enzymes such as that from S. cerevisiae, is involved
in forming the dimer–dimer interface. Insertions from dimeric
dPGMs exhibit sequence and conformational similarity to those
of tetrameric enzymes, illustrating that relatively small sequence
differences must define quaternary state. Indeed a single mutation
at this interface in S. cerevisiae dPGM dramatically destabilized
the tetrameric state without affecting kinetic properties [88].
Sequence differences between dimeric E. coli dPGM and the
tetrameric S. cerevisiae enzyme in this region have been analysed
[89].
In branch 2, the comparison of the E. coli glucose-1phosphatase structure with SixA again shows insertions after
β1 and β3, the former once again packing against the latter
(Figure 4e). Importantly, however, the β3–α3 insertion is very
large with approx. 100 residues. This insertion bears no obvious
sequence similarity to the branch 1 insertions at the same position,
and is composed almost entirely of α-helices. The E. coli glucose1-phosphatase structure contains other insertions far from the
catalytic site and a long C-terminal region, again apparently
unrelated to comparable regions in branch 1 of the superfamily.
The C-terminal extension forms a short region of antiparallel βsheet with β6. In E. coli phytase, the β3–α3 insertion is even
longer, at 140 residues almost as long as SixA itself (Figure 4f).
Finally, and remarkably, the A. fumigatus phytase structure differs
from the two branch 2 structures mentioned above in having
a large N-terminal extension, of largely irregular secondary
structure, which inserts into the back of the substrate-binding site,
opposite the entrance (Figure 4g), resulting in the availability of
Gln27 and Tyr28 for substrate interaction (see below).
The superimposed structures reveal that, quite surprisingly, the
entrances to the catalytic sites lie on opposite faces of the protein
in the two branches. Thus Figures 4(b)–4(d) show three branch 1
enzymes where substrate enters from the right past the C-terminal
regions (red) with important roles in allowing and responding
to substrate binding. The three branch 2 structures shown in
Figures 4(e)–4(g) contain catalytic sites accessed from the left.
This unusual situation has presumably been made possible by the
positioning of most of the catalytic apparatus after the central two
c The Authors Journal compilation c 2008 Biochemical Society
strands, β1 and β4, of the core β-sheet. In this way, additions to
the fold can form a substrate-binding site facing in either direction.
It is tempting to consider the simple SixA structure, with few
short loops between the regular secondary structure elements of
the core (Figure 4a), as resembling the ancestor of all present-day
members of the superfamily. By this hypothesis, evolution added
insertions to the basic scaffold to produce structures containing cavities suitable for binding of small-molecule substrates.
The absolute lack of sequence or structural similarity between the
respective insertions in branch 1 and branch 2 structures (Figure 4)
strongly suggests that this decoration of a basic fold happened
twice independently. Furthermore, the parallel processes of
elaboration on the basic fold began early, since both branches
are widely represented in both bacteria and eukaryotes.
CATALYSIS
Structural determinants
Catalytic activity in the superfamily centres on phosphorylation
and dephosphorylation of a histidine residue that follows the first
β-strand of the fold (His8 in E. coli SixA) (see Figures 5 and
6). A scheme for this catalytic cycle is shown in Figure 5.
The in-line transfer [90] of the phospho group from substrate
to enzyme occurs with the aid of several residues, forming a
‘phosphate pocket’, that hydrogen bond to the phospho group
before, during and after transfer. These include a pair of
flanking arginine residues, positions 7 and 55 in E. coli SixA
numbering, and the other histidine, His108 , which are completely
conserved in known active members of the superfamily (see,
e.g., Figure 3). Interestingly, the alignment also reveals a few
other residues that are highly conserved between otherwise
very diverse proteins (see also Figure 6). The most obvious
of these is Gly9 which forms part of the well-known and
characteristic RHG motif. Gly9 packs closely against the protein
core and is well-conserved as a glycine or occasionally an
alanine residue. The structure of E. coli glucose-1-phosphatase
(PDB code 1NT4 [91]) captures a very rare example of an
asparagine residue at this position and shows how this larger
side chain leads to structural displacements elsewhere in the fold.
The carbonyl of Gly9 makes an important hydrogen bond with the
phosphorylable histidine side chain holding it in an appropriate
conformation for catalysis (Figure 6). The backbone nitrogen of
Gly9 makes a further hydrogen bond with the side chain of Thr59
(SixA numbering) which is > 95 % conserved as threonine or
serine in both branches of the superfamily. The alignment also
reveals a L[S/T]XXG motif in the region between β1 and β2. As
shown in Figure 6, the glycine residue (numbered 27 in SixA) is
conserved for steric reasons; a larger side chain here would disturb
the catalytic apparatus. The leucine side chain contacts several
key residues (His8 , Gly9 and Arg55 ), while the threonine/serine
residue stabilizes the local structure through hydrogen bonds
to both the backbone of the neighbour of Gly9 (not sequenceconserved) and the backbone of Gly27 . Taken as a whole, this
hotspot of conservation in the second layer behind the catalytic
site illustrates how important maintenance of catalytic histidine
conformation is for activity.
Aside from the two histidine and two arginine residues
mentioned above, other interactions with the phospho group (the
PP residue in Figure 5) vary to a surprising extent (see Figure 3).
In most of branch 1, Asn16 (E. coli dPGM numbering) provides a
hydrogen bond. In the M. tuberculosis phosphatase [86], a serine
residue is present at this position and may substitute functionally,
although with the available crystal structure, and its nonphysiological ligands, it is hard to say. Any Asn16 -like interaction
is clearly absent in SixA, however, due to a local conformational
Structure and function in the histidine phosphatase superfamily
Figure 3
341
Structure-based sequence alignment of selected histidine phosphatase superfamily members
The Figure was generated by post-processing of a MUSTANG [83] structure alignment with STACCATO [84]. The structures shown are PDB codes 1UJC, E. coli SixA [85]; 1H2E, G. stearothermophilus
PhoE [92]; 1K6M, human liver F26BPase [46]; 1E58, E. coli dPGM [89]; 1NT4, E. coli glucose-1-phosphatase [91]; 1DKQ, E. coli phytase [103]; 1QWO, A. fumigatus phytase [93]. The secondary
structures of the smallest (1UJC) and largest (1QWO) proteins are shown below the alignment with β-strands of the core β-sheet (six in 1UJC, seven in 1QWO) and selected helices numbered. The
alignment is shaded blue according to sequence conservation with certain sets of residues (see text for details) picked out as follows: white on red, conserved catalytic core (see also Figure 6); black
on green, proton donors; black on yellow, additional members of the ‘phosphate pocket’; black on purple, substrate binding residues. Boxed blue residues are non-binding, but conserved positions
lying near the catalytic histidine (see Figure 6 and text). Note that the alanine mutations present at position 17 of 1DKQ and position 18 of 1NT4 have been replaced by the naturally occurring
histidine. The Figure was produced using JALVIEW [153].
c The Authors Journal compilation c 2008 Biochemical Society
342
Figure 4
D. J. Rigden
Cross-eyed stereo cartoons of structures of representative histidine phosphatase superfamily members
Structures are shown as cartoons and are coloured grey except for orange, N-terminal region; red, C-terminal tail; magenta, insertion in the β3–α3 region; blue, insertion in the β1–α1 region; and
turquoise, oligomeric dPGM-specific insertion (d). The structures are: (a) PDB code 1UJC, E. coli SixA [85]; (b) PDB code 1H2E, G. stearothermophilus PhoE [92]; (c) PDB code 1K6M, human
liver F26BPase [46]; (d) PDB code 1E58, E. coli dPGM [89]; (e) PDB code 1NT4, E. coli glucose-1-phosphatase [91]; (f) PDB code 1DKQ, E. coli phytase [103]; (g) PDB code 1QWO, A. fumigatus
phytase [93]. The phosphorylable histidine residue of SixA is shown as green spheres to localize the catalytic site.
difference [85]. An additional interaction with the phospho group
is provided by Gln22 in PhoE [92]. In SixA, allowing for a
small conformational change on substrate binding, Arg21 is well
positioned to interact with a phospho group [85], as predicted
[53]. The same modelling predicted that a different residue, Arg64
would fulfil the corresponding role in the Ais family [53]. Recent
modelling suggests that a further lineage in the superfamily, the
c The Authors Journal compilation c 2008 Biochemical Society
EPP/Sts-1 family, has independently evolved an arginine residue
at this position [20]. In branch 2 of the family, the phosphorylated
structure of A. fumigatus phytase (PDB code 1QWO; [93]) reveals
that the pair of arginine residues and second histidine residue are
supplemented by Arg62 which is positioned similarly to Asn16 in
E. coli dPGM [93]. The same interactions are predicted to hold
for all branch 2 enzymes of presently known structure.
Structure and function in the histidine phosphatase superfamily
Figure 5
343
Catalytic mechanism of the histidine phosphatase superfamily
The essentially invariant four residues of the catalytic core (see also Figures 3 and 6) are shown numbered as in E. coli SixA. His8 is phosphorylated during the course of the reaction. The other three
residues interact electrostatically with the phospho group before, during and after its transfer and form most or all of the ‘phosphate pocket’. Additional neutral or positive residues, represented as
PP in the diagram, may also contribute to the ‘phosphate pocket’ by hydrogen-bonding to the phospho group (see also Figure 3). The proton donor, an aspartate or glutamate residue whose position
varies in different families (Figure 3), is shown as PD.
Figure 6
The near-superimposable conserved catalytic cores of E. coli SixA and A. fumigatus phytase
The cores are shown in green and nearby conserved residues in purple for E. coli SixA (left; 156 residues; PDB code 1UJC [85]) and A. fumigatus phytase (right; 442 residues; PDB code 1QWO
[93]) which share 12 % sequence identity overall. Broken lines represent hydrogen bonds. Selected β-strands are drawn and labelled. The tungstate ion binding to the ‘phosphate pocket’ of SixA
is drawn as ball-and-stick in both panels. The Figure was generated using PyMOL (DeLano Scientific; http://www.pymol.org). A three-dimensional interactive version of this Figure can be seen at
http://www.BiochemJ.org/bj/409/0333/bj4090333add.htm.
For the most studied members of the superfamily, dPGM,
F26BPase and AP, there have been many mutagenesis studies
supporting the roles of the key catalytic residues (see, e.g.,
[88,94–98]). Typically, loss of these residues results in mutant
proteins with no or much reduced catalytic activity, but there
is an important exception that should be mentioned. When the
phosphorylable His256 of rat testis F26BPase was replaced with
an alanine residue, a highly unexpected 17 % of catalytic activity
remained [99], probably due to water taking over the nucleophilic
role [100]. This result argues that caution should be exercised
in inferring lack of catalytic activity for homologous sequences
lacking elements of the otherwise conserved catalytic machinery.
An important component of the catalytic scheme (Figure 5)
is the proton donor, required to donate a proton to the leaving
group as the substrate transfers its phospho group to the enzyme.
The only exception, not requiring proton donation, would be
priming of dPGM with 1,3-BPG, involving transfer of the acyl 1phosphate [4]. In most branch 1 members, a conserved glutamate
residue, Glu88 (E. coli dPGM numbering), functions as proton
donor [101]. It was therefore a surprise when it was realized
that the lineages having a minimal fold lack this residue [92]
and are predicted to use differently placed aspartate residues for
this function. The crystal structure of SixA [85] suggests that
the predicted conserved Asp18 is indeed likely to act as proton
donor. In branch 2, early mutagenesis data clearly pointed to
Asp304 (E. coli phytase numbering) as the proton donor [102],
and crystal structures show it to be well positioned with respect
to the rest of the catalytic apparatus. The side chain of Asp304 ,
c The Authors Journal compilation c 2008 Biochemical Society
344
D. J. Rigden
although contributed by a different part of the sequence, occupies
approximately the same position in superimposed structures as
Glu88 in E. coli dPGM. Asp304 as proton donor remains the
consensus view, but it has been argued that the equivalent to
the preceding histidine residue may, under certain circumstances,
act as a secondary proton donor, at least in glucose-1-phosphatase
[91]. This would require movement of the histidine side chain
from the crystallographically observed position [91]. It is worth
noting that such movement would be harder in branch 1 enzymes
since proton donor Glu88 , or non-catalytic Glu74 in SixA, packs
against the histidine side chain. A further suggestion is that
Asp304 and equivalents simply facilitate an intramolecular proton
movement on the substrate [71]. During the hydrolysis of the
phosphoenzyme structure, the same acidic residues activate bound
water molecules for nucleophilic attack. Crystal structures have
revealed suitably positioned water molecules in, for example, E.
coli dPGM [89] and E. coli phytase [103] complex structures.
When proton donors are highlighted on a sequence alignment
(Figure 3), it becomes clear that, within the superfamily, they can
be placed either shortly after strands β1 (SixA), β3 (dPGMs etc.)
or β4 (branch 2).
The anomalous mutases
Most of the superfamily are simple phosphatases carrying out the
scheme in Figure 5. However, there are anomalous members in the
form of the mutases: dPGMs and BPGMs. Each of these catalyses
three reactions (Table 1), but at very different relative rates [4].
Of these, the phosphatase reaction conforms to the scheme in
Figure 5. The ‘priming’ of dPGMs, i.e. their phosphorylation by
2,3-BPG or 1,3-BPG on the phosphorylable histidine residue, can
be considered as an incomplete phosphatase reaction. The ‘synthase’ reaction, the production of 2,3-BPG from 1,3-BPG, the
predominant activity of BPGM, would first involve the transfer
of the 1-phosphate of 1,3-BPG to enzyme. After reorientation of
the resulting 3PGA in the catalytic site, the phospho group
attaches to the 2-position and product 2,3-BPG is released from
the enzyme. The mutase reaction, catalysed at the highest rate by
dPGMs, involves binding of one phosphoglycerate form, either
3PGA or 2PGA, to the already phosphorylated (‘primed’) enzyme
and transfer of phospho group to the empty 2- or 3-position.
Intermediate 2,3-BPG then reorients within the catalytic site,
transferring the other phospho group to the enzyme. The primed
form of the enzyme is thereby restored and phosphoglycerate is
released. The distinguishing characteristic of mutases compared
with phosphatases is thus the ability to maintain intermediates
bound while allowing their reorientation with the active site. In the
case of dPGMs, a plausible mechanism for this reorientation has
been proposed [104]. Differences between dPGMs and BPGMs
relate first to the necessity of BPGMs to release 2,3-BPG, whereas
for dPGMs, it is the obligatory intermediate, and secondly to the
requirement of dPGMs to stabilize the phosphorylated form of
the enzyme and protect it from hydrolysis.
The C-terminal tail has long been implicated in stabilizing the
phosphoenzyme form of dPGM since the naturally tailfree dPGM from S. pombe and a proteolysed form of S. cerevisiae
dPGM lacking seven C-terminal residues, both have lower mutase
and higher phosphatase reaction rates when compared with
dPGMs having the tails [105]. Interpretation of these data was
complicated by the lack of interpretable density for the tail in
all S. cerevisiae structures. The position of the tail was only
seen on the determination of a structure of E. coli dPGM in
its phosphorylated form, strongly linking ordering of the tail and
phosphorylation state [89]. The earlier hypothesis that the tail
helped to exclude water from the catalytic site, and thereby avoid
c The Authors Journal compilation c 2008 Biochemical Society
hydrolysis of the phosphoenzyme [4], was rendered unlikely by
the presence of ordered water molecules in the catalytic site
of the phosphoenzyme structure [89]. Instead, the C-terminal
tail appears to provide additional stabilizing interactions with
the phosphorylated state, through interactions of tail residues
with Asn19 . This residue, and Asn16 , which forms two hydrogen
bonds with the phosphohistidine residue, lie on a loop whose
conformation varies according to phosphorylation state [106].
Interestingly, PhoE complex structures indicate that, although Cterminal tail structuring similarly accompanies modification of the
phosphorylable histidine residue, a different mechanism involving
Arg9 and the C-terminal carboxy group applies [92].
For dPGMs, there are no reliable structures of complexes
with substrates since there are reasons to doubt the validity
of the reported 3PGA complex [107], as discussed in [106].
Nevertheless, bound sulfate and vanadate seen in other structures
have offered clues as to likely substrate-binding residues
[104,106]. The binding of 2PGA and 3PGA (to phosphorylated
enzyme) and 2,3-BPG in two different orientations (as retained
intermediate in the mutase reaction, bound to unphosphorylated
enzyme) was modelled in E. coli dPGM [106]. The prediction
was that backbone amide groups of Thr22 and Gly23 bind the
carboxy group in all cases, and that side chains of Arg89 , Tyr91 ,
Arg115 , Arg116 and Asn185 bind the distal phospho group. For
BPGM, a remarkable set of complex structures, after various
periods of soaking in ligands, have recently provided a detailed
molecular picture of 2,3-BPG binding and transfer of its phospho
group to the enzyme [90]. The dPGM predictions are broadly
confirmed, suggesting that subtle differences are responsible for
the different spectra of activities of dPGM and BPGM. Different
explanations have been suggested, frequently involving Gly14
(human BPGM numbering; serine in dPGMs) and Ser24 (glycine
in dPGMs) [106,109]. Misleadingly, the first BPGM structure
[109] showed structural differences from dPGMs that turned out
not to be present in its ligated form [90]. Now that a direct
comparison of phosphorylated dPGM and BPGM can be made, it
is clear that neither residue obviously influences activity directly.
However, a noted restriction of catalytic size volume [109], in
part relating to differences in the C-terminal tail region, is still
apparent in BPGM compared with dPGM. This could easily
impede the reorientation of 2,3-BPG within the catalytic site that
lies at the heart of the mutase reaction. Additionally, although the
sets of substrate-binding residues (seen in BPGM, modelled in
dPGM) are very similar, a structural superposition shows many
conformational differences in their side chains that could relate
to different affinities for 1,3-BPG or 2,3-BPG between BPGMs
and dPGMs. One particularly interesting position is residue 100
(BPGM numbering), occupied by a conserved arginine residue in
BPGMs and by a conserved lysine residue in dPGMs. Deciphering
subtle differences in ligand affinities between the two families will
probably require the determination of a crystal structure of one in
complex with 1,3-BPG.
In passing, it is intriguing to note that the present superfamily is
not the only example of evolutionarily related phosphatases and
mutases, other examples being the cofactor-independent phosphoglycerate mutases, one of whose domains is homologous with
alkaline phosphatases [110] and β-phosphoglucomutase, closely
related to the phosphatase branch of the HAD (haloalkanoic acid
dehalogenase) enzyme superfamily [111]. In all cases, transient
phosphorylation of the enzyme is involved in catalysis.
Substrate binding in phosphatases
Information regarding which residues are responsible for substrate
binding in other members of the superfamily is not available for
Structure and function in the histidine phosphatase superfamily
all structures; indeed the substrate(s) of some enzymes of known
structure remain unknown [112]. In the case of F26BPase, the
crystal structure of the complex with fructose 6-phosphate and
phosphate [100], as well as mutagenesis data [113], implicate the
following set of residues in substrate binding: Ile267 , Tyr336 , Arg350 ,
Lys354 , Tyr365 , Gln391 and Arg395 (numbering for rat testis enzyme).
For PhoE, no complex is available with natural ligand, but its
preferred substrate, α-naphthyl phosphate, can be convincingly
docked into its notably hydrophobic active site, suggesting that
Met21 , Trp109 , Val153 , Leu86 and Phe108 are all substrate-binding
residues [114]. The crystal structure of E. coli SixA alone allowed
for the prediction of a tentative model of the complex with
substrate ArcB Hpt domain [85]. The model implicated SixA
residues Asn26 , Glu30 and Lys156 in substrate binding.
In branch 2 of the superfamily, complexes with inhibitors
(vanadate, tartrate, tungstate) are again more common than
substrate complexes. Nevertheless, substrate complex crystal
structures are available for both E. coli enzymes, the phytase and
the glucose-1-phosphatase [91]. The principal phytate-binding
residues in the former are Thr305 , Arg267 and Lys24 (there are
other interactions via water). In glucose-1-phosphatase, Glu196
and Tyr247 (via water) bind glucose-1-phosphate, whereas the
specificity of the enzyme for the 3-position of its alternative
substrate phytate is defined by the gating residue Leu24 [91].
The mode of interaction of phytate with A. fumigatus phytase
can be predicted based on the E. coli phytase complex [115]. In
this position, phytate interacts with the catalytic core plus Gln27 ,
Tyr28 , Lys68 , His189 , Lys278 , Tyr282 and Asn340 .
For future efforts in genome annotation in the superfamily, it
is important to know which regions of the enzymes are likely to
contribute to core catalysis and/or the determination of substrate
specificity. As mentioned above, the conserved catalytic core is
contributed by residues lying in the regions after strands β1, β2
and β4 (see Figures 3 and 6), whereas proton donors map to
the portions following β1, β3 and β4 (Figure 3). These are the
basic components for catalysis (although the surprising activity
of the H256A F26BPase mutant must be remembered). To these
may be added additional ‘phosphate pocket’ interactions, varying
between lineages, but, so far, always contributed by residues
lying in the β1–β2 loop (Figure 3). The pattern for specificitydetermining residues, binding the remainder of the substrate, is
very different. These are to be found scattered throughout the
sequence alignment, clustered in the loops following β1, β3
and β4, but also at various points in the large and very diverse
insertions between β3 and β4, in the N-terminus of A. fumigatus
phytase [93] and at the very C-terminus of SixA [85].
CONCLUSIONS
Although large-scale sequencing projects continue to discover
novel protein families awaiting function elucidation, function
predictions within superfamilies are equally valuable. The
histidine phosphatase superfamily, although possessing an everlengthening list of known functions, undoubtedly harbours many
novel activities awaiting discovery. Sequence and structure
analyses paint a clear picture of evolution of two deep-rooted
branches, the first of them, two thirds comprising bacterial
proteins, usually intracellular and of widely varied functions.
The second branch, predominantly eukaryotic and extracellular,
contains some well-known enzymes such as phytase and
other members, the APs, of partial or inconclusive functional
annotation. Humans contain several proteins from both branches,
many with known or potential medical importance. Phosphatases
in parasites may also have therapeutic value, whereas others have
(potentially) important agricultural applications.
345
After years of sporadic progress, the number of structures of histidine phosphatases is rising rapidly, allowing for a review of the
level of our understanding of the structure–function relationship
of the superfamily. All histidine phosphatase structures contain
a conserved catalytic core, and conserved ancillary residues
involved in maintaining the correct orientation of the core, but
vary dramatically in their proton donor and in substrate-binding
residues. This structural variability of specificity determinants
renders harder still function annotation in the superfamily and suggests that structural knowledge will be important, both for reliable
annotation of new sequences with known activities and for prediction of novel activities, prior to experimental demonstration. Potentially, both modelling (see, e.g., [20,53,116]) and further crystal
structures could be used for structure-based function predictions.
In summary, with many families and even structures
of unknown function, the histidine phosphatase superfamily
illustrates well the problems that have led experimentalists to
develop large-scale function screens [117], crystallographers to
suggest a structural genomics approach with superfamilies [118]
and bioinformaticians to seek novel structure-based function
assignment methods (see, e.g. [119]). An investigation of the
histidine phosphatase superfamily by any or all of these methods
would certainly yield many new insights.
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