Biochem. J. (2012) 442, 733–742 (Printed in Great Britain)
733
doi:10.1042/BJ20111443
2-Carboxy-D-arabinitol 1-phosphate (CA1P) phosphatase: evidence for
a wider role in plant Rubisco regulation
Paul John ANDRALOJC*1 , Pippa J. MADGWICK*, Yong TAO†2 , Alfred KEYS*, Jane L. WARD*, Michael H. BEALE*,
Jane E. LOVELAND*3 , Phil J. JACKSON‡, Antony C. WILLIS§, Steven GUTTERIDGE† and Martin A.J. PARRY*1
*Department of Plant Sciences, Rothamsted Research, Harpenden AL5 2JQ, U.K., †DuPont Stine-Haskell Research Center, Newark, DE 19714, U.S.A., ‡Chemical and Process
Engineering, University of Sheffield, Sheffield S3 7RD, U.K., and §MRC Immunochemistry Unit, University of Oxford, Oxford OX1 3QU, U.K.
The genes for CA1Pase (2-carboxy-D-arabinitol-1-bisphosphate
phosphatase) from French bean, wheat, Arabidopsis and
tobacco were identified and cloned. The deduced protein
sequence included an N-terminal motif identical with the
PGM (phosphoglycerate mutase) active site sequence [LIVM]x-R-H-G-[EQ]-x-x-[WN]. The corresponding gene from wheat
coded for an enzyme with the properties published for
CA1Pase. The expressed protein lacked PGM activity but
rapidly dephosphorylated 2,3-DPG (2,3-diphosphoglycerate) to
2-phosphoglycerate. DTT (dithiothreitol) activation and GSSG
inactivation of this enzyme was pH-sensitive, the greatest
difference being apparent at pH 8. The presence of the expressed
protein during in vitro measurement of Rubisco (ribulose1,5-bisphosphate carboxylase/oxygenase) activity prevented a
progressive decline in Rubisco turnover. This was due to the removal of an inhibitory bisphosphate that was present in the
RuBP (ribulose-1,5-bisphosphate) preparation, and was found
to be PDBP (D-glycero-2,3-pentodiulose-1,5-bisphosphate). The
substrate specificity of the expressed protein indicates a role for
CA1Pase in the removal of ‘misfire’ products of Rubisco.
INTRODUCTION
aestivum), Arabidopsis (Arabidopsis thaliana) and maize (Zea
mays) do not [7,8]. CA1P-inhibited Rubisco becomes reactivated
in an ensuing period of illumination by the combined action of
two light-activated, stromal enzymes: Rubisco activase, which
promotes the release of CA1P from the catalytic site of Rubisco,
and CA1Pase (CA1P phosphatase), which removes the phosphate
group of CA1P yielding Pi and CA (2-carboxy-D-arabinitol),
which is non-inhibitory. CA1P is made by the phosphorylation of
CA in low light or darkness [9,10], whereas CA itself is derived
from chloroplastic FBP (fructose-1,6-bisphosphate) [11].
Rubisco activity in vitro is seen to decline progressively as a
function of time and this phenomenon has been termed ‘fallover’.
It is caused by the accumulation of ‘misfire’ products of catalysis,
which remain tightly bound to the catalytic site on which they were
formed. The most important of the misfire inhibitors is likely to be
PDBP (D-glycero-2,3-pentodiulose-1,5-bisphosphate) (Figure 1),
which has also been shown to contaminate commercial and aged
preparations of RuBP [12,13]. Another by-product of Rubisco
turnover is XuBP (xylulose-1,5-bisphosphate) (Figure 1), which
is formed by the mis-protonation of the enediol derivative of RuBP
[14,15]. XuBP is a Rubisco substrate analogous to RuBP, but a
very poor one, since the rate of carboxylation of XuBP is orders
of magnitude lower than RuBP [16]. Thus XuBP is a competitive
inhibitor with respect to RuBP. Furthermore, RuBP and XuBP are
both inhibitory when bound to inactive (decarbamylated) Rubisco
[3,15].
The present study describes for the first time the identification
and cloning of the gene for CA1Pase and shows that the product
The Calvin cycle enzyme catalysing the reaction between CO2 and
acceptor molecule RuBP (ribulose-1,5-bisphosphate) (Figure 1) is
Rubisco (ribulose-1,5-bisphosphate carboxylase/oxygenase; EC
4.1.1.39). Rubisco also catalyses the reaction between O2 and
RuBP, initiating photorespiration. The activity of Rubisco is
regulated in vivo in several different ways. Long-term control
is achieved by altering the abundance of the protein through
gene expression and/or protein degradation. Short-term reversible
regulation of catalytic activity also occurs and involves active site
carbamylation, followed by the binding of an Mg2 + ion [1]. Lightdependent increases in stromal Mg2 + , pH [2] and Rubisco activase
activity all promote Rubisco activation and the converse is also
true [3].
Another strategy for regulation involves a tight binding
Rubisco inhibitor, synthesized under conditions of low light or
darkness [4]. This compound, CA1P (2-carboxy-D-arabinitol 1bisphosphate) is a naturally occurring, stable transition state
analogue of the carboxylase (CO2 -fixing) activity of Rubisco
(Figure 1) and binds exclusively to activated Rubisco [5,6].
Although CA1P is likely to be present in most plants [7], in many
plant species it does not accumulate sufficiently to have a significant effect on Rubisco activity. Thus French bean (Phaseolus
vulgaris), tobacco (Nicotiana tabacum), rice (Oryza sativa), soya
bean (Glycine max) and potato (Solanum tuberosum) accumulate
substantial amounts of CA1P (equivalent to 25–100 % of the
available Rubisco catalytic sites), whereas wheat (Triticum
Key words: 2-carboxyarabinitol 1-phosphate (CA1P), D-glycero2,3-pentodiulose-1,5-bisphosphate (PDBP) phosphatase, ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco).
Abbreviations used: BTP, Bis-Tris propane; CA, 2-carboxy-D-arabinitol; CA1P, CA 1-phosphate; CA1Pase, CA1P phosphatase; CABP, CA 1,5bisphosphate; CRBP, 2-carboxy-D-ribitol-1,5-bisphosphate; 2,3-DPG, 2,3-diphosphoglycerate; DTT, dithiothreitol; ESI, electrospray ionization; FBP,
fructose-1,6-bisphosphate; FBPase, fructose bisphosphatase; ORF, open reading frame; MS/MS, tandem MS; PDBP, D-glycero-2,3-pentodiulose-1,5bisphosphate; PFK, phosphofructokinase; PGM, phosphoglycerate mutase; Q–TOF, quadrupole–time-of-flight; RACE, rapid amplification of cDNA ends;
Rubisco, ribulose-1,5-bisphosphate carboxylase/oxygenase; RuBP, ribulose-1,5-bisphosphate; TEA, triethanolamine; TSP-d4 , 3-(trimethylsilyl)-2,2 ,3,3 tetradeuteropropionic acid; XuBP, xylulose-1,5-bisphosphate.
1
Correspondence may be addressed to either of these authors (email [email protected] or [email protected]).
2
Present address: Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, People’s Republic of China.
3
Present address: Wellcome Trust Sanger Institute, Hinxton CB10 1HH, U.K.
c The Authors Journal compilation c 2012 Biochemical Society
734
P. J. Andralojc and others
Figure 1 Structural formulae of naturally occurring substrates and
inhibitors of Rubisco: RuBP, CA1P, PDBP and XuBP
of its expression has properties which are consistent with those
described previously for CA1Pase purified from plant leaves. We
demonstrate that the redox regulation of its catalytic activity is pHdependent and that it has a substrate specificity which indicates a
role for CA1Pase in the removal of misfire products of catalysis.
EXPERIMENTAL
Identifying and expressing the coding sequence
CA1Pase was purified from fresh leaves of P. vulgaris L. (cv.
Tendergreen) as described previously [17]. In separate digestions,
the purified protein was treated with trypsin, chymotrypsin, V8
protease and CNBr and the resolved peptide fragments were
transferred to PVDF membranes for gas-phase peptide sequencing
(Procise® protein sequencing system; Applied Biosystems) using
a modified Edman chemistry cycle [18]. Protein database analysis
(BLASTP, National Centre for Biotechnology Information)
revealed that many of the bean peptides had distinct homologies
with co-factor dependent PGMs (phosphoglycerate mutases) and
so the full-length sequence of this gene was obtained from the
Arabidopsis genome (At5g22620). Degenerate oligonucleotides
were designed to bind to regions of the gene encoding two of
the bean peptides (TAEIIWG and QWQIDAENFIIDGHYPVR,
extending the sense and antisense strands of the gene respectively)
and to a highly conserved region in PGM (HGQSTWN, extending
the sense strand of the gene). These were used to amplify and clone
part of the gene, followed by 3 - and 5 -RACE (rapid amplification
of cDNA ends) to obtain the complete cDNA sequence for this
gene from bean and tobacco. Although these genes incorporated
a chloroplast transit peptide (as identified by ChloroP [19])
these were omitted from our expression studies. The amino acid
sequence of the enzyme was also used to identify a clone from
a DuPont cDNA collection that originated from a wheat library.
This clone contained the complete ORF (open reading frame)
of the enzyme from which the gene could be amplified with
primers that introduced useful restriction sites at the 5 and 3 ends.
Two primers, 5 -GATCACCATGGCGAAGAGGGTGGTTCT-3
and 5 -TGAGCTCGAGCCTTAAAACTCTTCACCA-3 , were
designed to introduce an Nco1 site coincident with the 5 -ATG of
the ORF and an Xho1 site adjacent to the 3 termination codon
respectively. The mature wheat gene was expressed in Escherichia
coli (strain BL21) using expression vector pBX3, replete with
T7 promoter region, His6 tag and multiple cloning sites. Our
studies utilized the His6 -tagged expression product of the wheat
homologue. All procedures, from the expression of the cloned
CA1Pase by E. coli in liquid culture through to enzyme isolation,
were by standard techniques. Ni2 + -NTA (Ni2 + -nitrilotriacetate)–
agarose (Qiagen) was used for affinity purification, after which
c The Authors Journal compilation c 2012 Biochemical Society
any contaminating proteins were removed by gel filtration
chromatography using a column (65 cm long × 1.6 cm diameter)
of Superdex-75 and column buffer 150 mM NaCl/50 mM BTP
(Bis-Tris propane), pH 7.0, running at 0.5 ml/min. Peak fractions
were pooled, divided into 0.2 ml aliquots, frozen in liquid nitrogen
and stored at − 80 ◦ C. Prior to enzyme assay, thawed aliquots
were spin-desalted through small columns (2 ml) of Sephadex
G-25 medium, pre-equilibrated with 100 mM BTP, pH 7.0, and
the concentration of the recovered protein (typically 1–2 mg/ml)
was determined by the method of Bradford [23] relative to a
BSA (Fraction V; A3059, Sigma) standard curve. The desalted
protein could be repeatedly snap-frozen and thawed, without loss
of catalytic activity. Dilution of desalted CA1Pase into buffers
containing 1 mg/ml BSA, ensured that the catalytic activity was
directly proportional to the dilution, to a concentration of less than
1 μg of CA1Pase/ml.
CA1Pase assay
CA1Pase activity was measured by the time-dependent
appearance of Pi [20]. Assay buffer contained 0.1 M BTP,
pH 7.0, and 1 mg/ml BSA with other components at the stated
concentrations. Prior to assay, desalted CA1Pase was diluted to
66 μg/ml in 0.1 M BTP, pH 7.0, and 1 mg/ml BSA to provide
a common stock, and used at the rate of 5 μl per 95 μl assay
mixture. CA1Pase was added to start each assay. 0, 30, 60 and
90 s later, 100 μl aliquots of the reaction solution were quenched
with 15 μl of ice-cold 2 M TCA (trichloroacetic acid). After
20 min at 0 ◦ C the precipitated protein was sedimented by
centrifugation (microfuge: 13 100 g, 10 min, 4 ◦ C) and 100 μl
of the supernatant transferred to a 96-well microtitre plate. Pi
standards (1–7 nmol) of the same volume and composition as the
quenched supernatants, were prepared in parallel. Then 200 μl of
0.44 % (w/v) ammonium molybdate in 1.6 M H2 SO4 was added,
followed 10 min later by 50 μl of 0.035 % (w/v) Malachite Green
in 0.35 % (w/v) poly(vinyl alcohol), with thorough mixing. After
60 min at room temperature (22 ◦ C) the A610 was determined
(SpectraMax 340PC; Molecular Devices). Pi release usually
varied linearly with time. If not, the assay was repeated with less
enzyme and/or shorter time intervals. In assays that included DTT
(dithiothreitol) a separate, parallel, set of Pi standards containing
an identical quantity of DTT was prepared as it was found to
affect colour development. The activities given are the mean and
S.D. of the initial activity, obtained from a series of time points,
performed simultaneously.
In reaction mixtures containing CA1P and a phosphatecontaining effector, CA1P phosphatase activity was deduced from
the time-dependent release of 14 C-labelled 2-carboxyarabinitol
([14 C]CA) from [14 C]CA1P [21]. The reaction, in a total volume of
100 μl, contained 0.1 M TEA (triethanolamine), pH 8.0, 1 mg/ml
BSA, 0.2 mM CA1P (102 kBq of [21 -14 C] CA1P/μmol of CA1P),
5 mM MgCl2 , 5 mM effector and 3.3 μg of CA1Pase. Before
assaying, desalted CA1Pase was diluted to 66 μg/ml in 0.1 M
TEA, pH 8.0, 1 mg BSA/ml and 10 mM DTT to provide a
common stock. This was kept at 25 ◦ C for 60 min to fully activate,
then promptly used at the rate of 5 μl per 95 μl assay mixture,
to start each assay. The [14 C]CA released was measured by liquid
scintillation counting after removal of remaining [14 C]CA1P
by anion exchange (Dowex-1). Each assay was performed in
triplicate and the results expressed as means +
− S.D.
Rubisco activity
Rubisco activity was determined in 500 μl of assay buffer
comprising 100 mM Bicine, pH 8.0, 20 mM MgCl2 , 10 mM
NaH[14 C]CO3 (18 kBq/μmol), 0.4 mM RuBP and 12.5 μg of
CA1P phosphatase and Rubisco regulation
activated Rubisco, at room temperature. When assaying the
inhibitory activity of HPLC-resolved fractions, Rubisco and 25 μl
of the fraction indicated were pre-incubated in assay buffer
(without RuBP) for 10 min, after which RuBP was added to
initiate the activity assay. When assessing the effect of CA1Pase
on RuBP carboxylation, 3.75 μg of CA1Pase was added to the
assay buffer 15 min before inclusion of Rubisco, which was
added to initiate the assay. Unless otherwise indicated, the assay
duration was 5 min. All assays were quenched with 100 μl of
10 M formic acid. The quenched samples were evaporated to
dryness, rehydrated in 400 μl of water, mixed with scintillation
cocktail and the acid-stable 14 C determined as above. Each time
course was performed in triplicate and the results expressed as
means +
− S.D.
HPLC
RuBP and phosphorylated derivatives of RuBP were resolved
by anion-exchange HPLC using a CarboPac PA1 analytical
(4 mm × 250 mm) and guard (4 mm × 50 mm) column in
conjunction with a Dionex DX500 chromatography system,
including an absorbance (AD20) and an electrochemical (ED40)
detector with gold working electrode and silver–silver chloride
reference electrode. Aqueous samples of 20–100 μl, from which
divalent metal cations had been removed, were injected following
column equilibration with 80 % (v/v) water, 20 % (v/v) 1 M
sodium acetate, pH 7.0, and a flow rate of 1 ml/min. After
injection, the proportion of 1 M sodium acetate in the aqueous
mobile phase (v/v) changed linearly as follows: 5–30 min, 20–
100 %; 30–32 min, 100–20 %; 35–40 min, 20–100 %; 45–48
min, 100–20 %; 60 min, next sample injection. (Inclusion of a
second sodium acetate gradient cycle, as indicated, was required to
fully regenerate the column.) Fractions were collected at 0.5 min
intervals from 15.0 min (retention time of earliest inhibitor,
CA1P) to 25.0 min [at least 0.5 min after elution of the strongly
retained synthetic inhibitor, CABP (2-carboxy-D-arabinitol 1,5bisphosphate)] and frozen in liquid nitrogen as soon as possible
after collection. The chromatograms shown are representative of
the replicate chromatograms which were performed. In addition,
a MonoQ (100 mm long × 10 mm diameter) column was used
to fractionate inhibitors derived from RuBP. A tertiary gradient
system was used, where solution A was pure water, solution B
was 0.5 M NaCl, and solution C was 50 mM Hepps, pH 8.0, and
50 mM sodium borate. The composition of the mobile phase (v/v)
was as follows: 0–0.1 min, 75 % solution A, 5 % solution B,
20 % solution C, changing linearly thereafter to 57.4 % solution
A, 22.6 % solution B and 20 % solution C by 90 min. This was
followed by column regeneration (0 % solution A, 80 % solution
B and 20 % solution C for 5 min) and then a return to the initial
conditions 15 min before the next injection. Injection volume was
1 ml, the flow rate was 2 ml/min, and peak detection was by
monitoring UV absorbance at the stated wavelength.
Rubisco purification
Wheat Rubisco was prepared essentially as described in [22].
Soluble protein concentrations for Rubisco and CA1Pase were
determined using the method of Bradford [23] with BSA standard
curves prepared in parallel.
1
H-NMR analysis of inhibitors
Sample (600 μl) containing ∼ 30–50 nmol of the quinoxaline
derivative was added to 50 μl of 2 H2 O and 20 μl of
735
H2 O/[2 H]methanol (80:20) containing 0.05 % (w/v) sodium
TSP-d4 [3-(trimethylsilyl)-2,2 ,3,3 -tetradeuteropropionic acid].
1
H-NMR was acquired at a temperature of 300 K on an Avance
600 spectrometer (Bruker Biospin) operating at 600.0528 MHz
using a 5 mm SEI probe. The residual 2 H2 O signal was suppressed
by pre-saturation and a relaxation delay of 5 s was employed.
Each spectrum consisted of 2048 scans of 64 000 data points
with a spectral width of 12 p.p.m. FIDs (free induction decays)
were automatically Fourier transformed after the application of
an exponential window function with a line broadening of 0.5 Hz.
1
H-NMR chemical shifts were referenced to the (CH3 )3 signal of
TSP-d4 at δ 0.00.
2
ESI (electrospray ionization)–MS of inhibitors
Sample (50 μl) containing 2.5–5 nmol of the quinoxaline
derivative diluted in 400 μl of water/methanol (4:1, v/v) was
infused into the spectrometer (Esquire 3000; Bruker Daltonics)
by flow injection using an Agilent 1100 series HPLC system with
degasser, quaternary pump and autosampler. The flow rate was
100 μl/min of 20 % (v/v) methanol in water. Mass spectra were
recorded from 1.7 to 4.2 min after the sample had entered the flow.
Spectra were recorded in both positive and negative ion mode on
the same sample via an alternating sequence of the two ionization
modes, each time recording the average of 25 scans. Spectra were
recorded over an m/z range of 50 –1000. The spectra were recorded
using Ion Charge Control with a maximum accumulation time of
40 ms for 20 000 (negative mode) or 50 000 (positive mode) ions.
The nebulizer pressure was 20 psi (1 psi = 6.9 kPa) and the dry
gas was at a flow rate of 6 litres/min at 350 ◦ C.
Accurate mass determination
Accurate mass determinations on ions identified in ESI–MS were
carried out on a Waters Q –TOF 1 (quadrupole–time-of-flight 1)
instrument. Then 100 μl of quinoxaline samples diluted in 900 μl
of water/methanol (4:1, v/v) was infused directly into the mass
spectrometer by flow injection using a syringe pump at flow rates
of 30 and 50 μl/min and desolvation temperatures of 180 and
150 ◦ C for positive ion and negative ion analysis respectively. The
source temperature was 100 ◦ C and the desolvation and nebulizer
gas flows were 400 and 25 litres/h respectively. Spectra were
recorded over an m/z range of 100 –1700 with an acquisition rate
of 2.4 s per scan. Accurate mass spectra were determined using
an external mass axis calibration and internal lock (correction)
mass. Mass axis calibration employed sodium iodide (ESI + ) or a
mixture of sugars (G1 – G9 , ESI − ).
Substrate synthesis
XuBP was synthesized from FBP and glycoaldehyde phosphate
[24] including subsequent product purification by Dowex-1
chromatography. Glycoaldehyde phosphate was synthesized
as described [25]. RuBP, CABP, CRBP (2-carboxy-D-ribitol1,5-bisphosphate) and CA1P were synthesized as described
previously [26] and CA was derived by alkaline phosphatase
treatment of CABP, followed by anion-exchange purification of
the product, using Dowex-1. An inhibitory derivative of RuBP,
PDBP, was synthesized by the method of Kane et al. [12] by
incubating pure RuBP (5 mM) with an oxygen-saturated solution
containing 0.1 M Hepps, pH 8.0, 40 mM boric acid, and 5 mM
CuSO4 , in a total volume of 1 ml for 2 h at room temperature,
followed by passage through a 0.5 ml bed volume of Chelex 100.
The yield of PDBP by this method did not exceed 5 % of the
original RuBP, as determined by a Rubisco inhibition assay.
c The Authors Journal compilation c 2012 Biochemical Society
736
Figure 2
P. J. Andralojc and others
Amino acid sequence of a putative PGM from Arabidopsis (At): At5g22620 (CA1P phosphatase)
This sequence has been aligned with homologous sequences from tobacco (Nicotiana tabacum, Nt; accession number HE610108), French bean (Phaseolus vulgaris, Pv; accession number HE603917)
and wheat (Triticum aestivum, Ta; accession number HE603918). The asterisk (∗) marks the first amino acid in the At mature polypeptide, as predicted by ChloroP. The # symbol marks the amino
acid to which the N-terminal histidine peptide MRGSHHHHHHSM was attached in the expressed protein from wheat (Ta). Backgrounds: dark grey, residues common to all sequences; light grey,
conservative substitutions and residues common to some sequences; white, dissimilar residues.
The resulting solution was either used immediately or snap-frozen
and stored at − 80 ◦ C.
Other materials
Sodium [14 C]bicarbonate was supplied by GE Healthcare. Dowex
(AG 50W-X8 and AG 1-X8) and Chelex resins were supplied
by Bio-Rad. All other reagents, including the ‘commercial’
preparation of RuBP referred to later, were supplied by Sigma
and were of analytical grade.
active site sequence. These were used to amplify and clone
part of the gene, followed by 3 - and 5 -RACE to obtain the
complete cDNA sequence for this gene from bean and tobacco.
The homologous gene from wheat was similarly identified and
cloned. The derived amino acid sequences for the homologous
enzymes from Arabidopsis, French bean, tobacco and wheat
are illustrated in Figure 2. The GenBank® accession numbers
for these sequences are HE603917 (French bean), HE610108
(tobacco) and HE603918 (wheat).
Establishing that the expressed gene is a CA1Pase
RESULTS
Cloning the CA1Pase gene
As previously reported, CA1Pase purified from P. vulgaris was
catalytically active and ran as a single band during SDS/PAGE
[26]. Attempts at N-terminal sequencing using the intact
enzyme were unsuccessful but a variety of protein fragmentation
techniques (see the Experimental section) followed by sequence
analysis by Edman degradation [18] provided internal amino acid
sequence data. Protein database analysis (BLASTP) revealed that
many of the bean peptides had distinct homologies with co-factor
dependent PGMs and so the full-length sequence of this gene was
obtained from the Arabidopsis genome (At5g22620). As detailed
in the Experimental section, degenerate oligonucleotides were
designed to bind (i) to regions of the gene encoding two of
the bean-derived peptides, and (ii) to a highly conserved PGM
c The Authors Journal compilation c 2012 Biochemical Society
Expression of a His6 -tagged wheat homologue revealed a protein
which migrated as a single band on SDS/PAGE with an
apparent mass of 56 kDa (Supplementary Figure S1 available at
http://www.BiochemJ.org/bj/442/bj4420733add.htm) and which
also had a retention time consistent with a monomeric protein
during size-exclusion HPLC (BioSep S4000, Phenomenex;
results not shown). This protein had CA1Pase activity (Figure 3)
which varied linearly with assay duration and enzyme abundance
(Supplementary Figure S1). The wheat homologue had a K m
−1
for CA1P of 10 +
− 1 μM and a V max of 4.2 +
− 0.1 μmol/min
per mg of protein (Figure 3). As expected for CA1Pases
[26–28] this enzyme also dephosphorylated the closely related
compounds CABP and CRBP at a similar rate to CA1P (Figure 3,
inset table). However, the enzyme had higher affinity for these
bisphosphates than for CA1P, with K m values of one-quarter and
one-half of that for CA1P respectively. Even when the reaction
CA1P phosphatase and Rubisco regulation
Figure 3
737
Substrate preference of wheat CA1Pase
Values of V max and K m as defined by the Michaelis–Menten equation were determined by an
iterative, non-linear regression algorithm (Enzfitter, Elsevier Biosoft). Substrate concentrations
ranged from 2.5 to 1000 μM, with assay duration of 15, 30, 60 and 120 s at each concentration,
at pH 7.0, and the release of Pi was determined spectrophotometrically. RuBP and XuBP were
found to decompose in the presence of the acidic molybdate reagent used in the determination
of free phosphate. However, HPLC analysis of RuBP and XuBP (conducted at neutral pH) after
incubation with CA1Pase showed no hydrolysis of either bisphosphate, even after 10 min in the
presence of 5-fold more CA1Pase than was used for the other potential substrates. Since they
were not substrates for this CA1Pase, K m determination was not applicable (n.a.) as indicated.
mixture was analysed by HPLC, we found no evidence for the
dephosphorylation of either RuBP or its analogue, XuBP, even
after prolonged incubation with the expressed enzyme, leading
us to conclude that neither of these compounds were substrates.
In spite of the enzyme being homologous with PGMs, there was
no activity, either as a phosphatase or as a mutase, towards 3phosphoglycerate. However, one class of PGMs with significant
homology with the CA1Pase genes identified in the present study
are co-factor 2,3-DPG (2,3-diphosphoglycerate)-dependent. We
found that the CA1Pase had considerable phosphatase activity
towards this metabolite, yielding 2-phosphogylcerate, albeit with
an affinity that was 20-fold lower than for CA1P (Figure 3, inset
table).
We investigated the effect of pH and redox modulation on the
activity of the expressed wheat CA1Pase (Figure 4A). Aliquots of
the CA1Pase preparation were pre-incubated for 60 min at 25 ◦ C
in the presence or absence of 10 mM DTT or 5 mM GSSG at
pH 6.0, 7.0 and 8.0 (Figure 4A) prior to assay initiation by the
addition of CA1P. All forms of the enzyme were most active at
pH 7, in agreement with reports for the pH optima of the tobacco
and bean enzymes, purified from leaf extracts [26,29]. The activity
of the expressed enzyme was only slightly increased by preincubation with DTT at pH 8.0, but not at pH 6.0 or 7.0. However,
the activity showed significant attenuation after pre-incubation
with GSSG, especially at pH 8.0, where the activity was 10-fold
lower than the DTT-treated counterpart (Figure 4A). In other
words, a greater degree of activity regulation was observed at
pH 8 than at pH 7, although the catalytic capacity of the reduced
enzyme was greatest at pH 7.
A variety of phosphorylated metabolites have been shown to
influence the rate of dephosphorylation of CA1P by CA1Pase
[26,28–30]. In agreement with these previous studies, ADP, PPi
(pyrophosphate), FBP, RuBP, 3-PGA (3-phosphoglyceric acid)
and 2-PG (2-phosphoglycolate) were all stimulatory, roughly
doubling the rate of CA appearance (Figure 4B). However, Pi
Figure 4 Redox regulation and the influence of physiologically relevant
metabolites on the activity of His6 -tagged wheat CA1Pase
(A) Redox regulation. The CA1Pase preparation was pre-incubated for 60 min at 25 ◦ C with
BTP/BSA buffer alone (white bars) or supplemented either with 10 mM DTT (grey bars) or 5 mM
GSSG (black bars) at pH 6.0, 7.0 and 8.0, then immediately assayed for CA1Pase activity by
measuring the release of Pi . (B) Effect of phosphorylated metabolites on the dephosphorylation of
[14 C]CA1P by CA1Pase. As described in the text, the effect of each metabolite (at a concentration
of 5 mM) was determined by the time-dependent evolution of [14 C]CA at pH 8.0, rather than by
the release of Pi . Results are means and S.D. for three replicate determinations.
was found to have little effect on activity, although it was found to
be inhibitory in the earlier studies. This discrepancy may be due
to differing sensitivities to Pi between homologous enzymes from
different species, as exemplified by the response to ATP, which is
stimulatory in P. vulgaris [28], but inhibitory in tobacco [29].
Effect of CA1Pase on Rubisco carboxylase activity
The presence of CA1Pase improved the linearity of the timedependent carboxylation of RuBP by Rubisco. In the absence
of CA1Pase (Figure 5A, 䊊) the production of acid-stable
14
C by Rubisco progressively declined, but in its presence
(Figure 5A, 䊉) the decline was far less pronounced. This
difference became more significant during the course of the
assay. Furthermore, this phenomenon was completely abolished
when the phosphatase was heat denatured prior to inclusion in
the assay (Figure 5B, ×), implying that the stimulation was
dependent on its catalytic activity. Analysis of the commercial
preparation of RuBP used in these assays by anion-exchange
HPLC with UV detection (Figure 5D) revealed a large RuBP
peak (18 min) followed by a smaller contaminating peak at
21.7 min (denoted by ‘?’). When RuBP synthesized in-house
(‘RRes’) was similarly resolved by HPLC, the corresponding
contaminant was virtually absent (Figure 5D). In parallel Rubisco
assays using both RuBP preparations, only the commercial
preparation showed significantly greater carboxylation in the
presence of the CA1Pase (Figure 5C, white compared with
light grey bars), while the carboxylation of RRes RuBP was
uniformly high (Figure 5C, mid compared with dark grey bars).
It therefore appeared that the CA1Pase-dependent enhancement
was connected to the contaminating peak in the commercial
c The Authors Journal compilation c 2012 Biochemical Society
738
Figure 5
P. J. Andralojc and others
Effect of CA1Pase on RuBP carboxylation by Rubisco
(A and B) Time-dependence of Rubisco carboxylase activity in the presence and absence of
wheat CA1Pase. (A) CA1Pase (3.75 μg, 䊉) or an equivalent volume of BTP/BSA buffer alone
(control, 䊊) were added to the assay buffer 15 min before the addition of 12.5 μg of activated
Rubisco, to initiate the assay. Assay duration varied between 0 and 5 min, as shown. (B) Similar
to (A) but highlighting the effect on Rubisco activity of pre-incubating RuBP with an identical
amount of heat-treated CA1Pase (×). Results in (A and B) are mean values for three replicate
determinations, with the error bars being obscured by the data symbols. (C) Comparing the
effect of pre-incubating a commercial (white and light grey) and a purer ‘RRes’ (mid and dark
grey) RuBP preparation with (light and dark grey) or without (white and mid grey) CA1Pase, on
their subsequent carboxylation by Rubisco. Results for 0, 5 and 10 min pre-incubations with and
without CA1Pase are shown. The duration of the subsequent carboxylase assays was 5 min. The
mean and S.D. for three replicate determinations are given. (D) HPLC analysis of a commercial
(upper trace) and of a purer ‘RRes’ (lower trace) RuBP preparation. HPLC resolution utilized
a CarboPac PA1 column in conjunction with a neutral acetate gradient with peak detection
at 280 nm. The same commercial preparation of RuBP (R0878, Sigma) was used throughout
(A–D). The purer ‘RRes’ RuBP (C and D) was synthesized at Rothamsted Research.
RuBP preparation. Furthermore, since the CA1Pase-dependent
enhancement of carboxylation of the purchased RuBP was only
very slightly increased by pre-incubating RuBP with CA1Pase
prior to assay initiation (Figure 5C) the phosphatase must exert
its effect relatively quickly.
The commercial RuBP preparation was pre-incubated
in the absence (Supplementary Figure S2A available at
http://www.BiochemJ.org/bj/442/bj4420733add.htm) or presence (Supplementary Figure S2B) of CA1Pase, followed
by ultrafiltration, HPLC analysis and fractionation. Fractions
collected from the emergence of RuBP onwards were assayed
for the presence of Rubisco inhibitors by measuring their
effect on Rubisco carboxylase activity. In the absence of
CA1Pase, two troughs of inhibition were detected (Figure S2A):
one immediately after the elution of RuBP (which itself had
a stimulatory effect) and another which coincided with an
absorbance peak at ∼ 21.7 min. In the presence of the CA1Pase,
both troughs of inhibition were diminished, as was the UV
absorbing peak at 21.7 min. It therefore seemed likely that two
distinct Rubisco inhibitors were present in the commercial RuBP
preparation, one of which absorbed UV light, which were both
diminished by CA1Pase treatment. The complete removal of
inhibitors by CA1Pase was not seen to occur. This may have
been due to the continued presence of RuBP, from which more
inhibitor may have been generated, following the removal of
CA1Pase prior to HPLC. An additional, spurious, contaminating
peak at 21 min was apparent in the commercial RuBP illustrated
in Supplementary Figure S2(A). However, this did not inhibit
Rubisco.
c The Authors Journal compilation c 2012 Biochemical Society
Figure 6
Detailed analysis of Cu2 + /O2 oxidized RuBP
(a and b) 10 μl of Cu2 + -treated RuBP, in a final volume of 100 μl was incubated with
BSA/BTP-HCl buffer, pH 7.0, alone (a) or supplemented with 6 μg of wheat CA1Pase (b).
After 20 min at room temperature, the samples were treated with excess Dowex-50 (H + -form),
filtered, and aliquots (50 μl) were resolved by anion-exchange HPLC at pH 7.0 (a and b). (c) An
aliquot (50 μl) of the sample described in (a) was resolved by HPLC and the eluate collected into
0.5 ml fractions and frozen immediately in liquid N2 . Then aliquots (5 μl) from each fraction were
pre-incubated with 12.5 μg of activated Rubisco for 5 min and the resulting Rubisco activity
determined. 100 % = 0.20 μmol of CO2 fixed/min per mg of protein. (d) Cu2 + -treated RuBP
(20 μl) was resolved as in (a), except peak detection was by pulsed amperometry, following
post-column addition of 0.3 M NaOH at 0.5 ml/min. (e) Aliquots (20 μl) of the oxidized RuBP
were again fractionated by HPLC and the (0.5 ml) fraction corresponding to the peak shown in
(a) was treated with excess Dowex-50 (H + -form) then snap-frozen. Then 100 μl of this was
analysed as in (d). (f) An aliquot (20 μl) of the oxidized RuBP was fractionated by HPLC and the
fractions containing the earlier eluting inhibitor (corresponding to the peak at 19 min in d) were
combined, treated with excess Dowex-50 (H + -form) then snap-frozen. Then 100 μl of this was
analysed as in (d). (g) As in (f), except the isolated earlier-eluting inhibitor was incubated with
wheat CA1Pase for 60 min at room temperature prior to HPLC analysis.
Properties of the inhibitory contaminants
When pure RuBP (synthesized in-house) was treated with Cu2 +
ions in the presence of dissolved oxygen, as described for the
formation of PDBP by Kane et al. [12], and then analysed by
anion-exchange HPLC, a predominant peak with a strong UV
absorbance and a retention time identical with that of the
UV absorbing inhibitory contaminant of the commercial RuBP
was detected (Figure 6a). This suggested that one of the inhibitory
contaminants removed by CA1Pase was PDBP, although UV
absorbance of this compound had not been reported previously.
An identical peak was also obtained when XuBP was treated
in the same way (results not shown). The size of this strongly
absorbing peak was diminished by prior exposure to CA1Pase
(Figure 6b). When the HPLC fractions from the untreated RuBP
derivative (shown in Figure 6a) were pre-incubated with Rubisco
and the resulting carboxylase activities determined (Figure 6c),
CA1P phosphatase and Rubisco regulation
Figure 7 Structural formulae of the 2,3-substituted quinoxaline of PDBP as
a free acid (M r 380) and as a monosodium salt (M r 402)
two roughly equal troughs of inhibition were apparent, indicative
of two distinct bisphosphate inhibitors, with retention times
identical with those identified in the commercial preparation of
RuBP (Supplementary Figure S2A). Clearly, the earlier eluting
inhibitor lacked the prominent UV absorbance of the later eluting
component, although fractions from both were found to contain
similar amounts of inhibitory activity. When the Cu2 + -treated
RuBP was analysed by HPLC and peak elution monitored
electrochemically (by integrated amperometry), it was seen to
contain three prominent peaks (Figure 6d) with the earliest
eluting of these having the retention time of RuBP, followed
by two peaks whose positions coincided with the two troughs
of inhibition. When HPLC fractions containing the inhibitory,
UV absorbing, component collected between 21 and 22 min
(as seen in Figure 6a) were subjected to a second, identical,
round of HPLC, they were found to elute as two predominant
peaks, which coincided with the two peaks of inhibitory activity
(Figure 6e). Conversely, when HPLC fractions containing the
earlier-eluting, non-UV absorbing, inhibitor was subjected to a
second, identical, round of HPLC, they were also found to yield
peaks coinciding with both inhibitory components as well as
RuBP (Figure 6f). Thus the two inhibitory compounds appear to
be interconvertible. Furthermore, when the HPLC-fractionated,
earlier eluting, inhibitory component (Figure 6f) was treated with
CA1Pase before re-analysis by HPLC, both of the peaks which
coincided with the two inhibitors were greatly diminished, but not
the peak corresponding to RuBP (Figure 6g). Thus the amount of
both inhibitors is diminished by CA1Pase.
Identifying the inhibitory contaminants
There was insufficient inhibitory material in our commercial
RuBP preparation to permit spectroscopic identification.
However, the identical properties of these inhibitory contaminants
with those of the inhibitors formed by the Cu2 + -catalysed
oxidation of RuBP implied that they were identical. We therefore
attempted to elucidate the identity of the larger amounts of
inhibitor present in preparations of Cu2 + -oxidized RuBP. PDBP
has been reported as being extremely labile [12] and so we
attempted to derivatize the inhibitors with o-phenylenediamine
as soon as they emerged during HPLC fractionation. As shown
previously [12,14] any vicinal dicarbonyl compounds present
(such as those in PDBP) would be converted into relatively
stable 2,3-substituted quinoxalines (Figure 7) in the presence of
o-phenylenediamine. However, whereas the resolving power of
the CarboPac PA1 column (as demonstrated by Figures 5 and
6, and Supplementary Figure S2) was excellent, the associated
requirement for a sodium acetate gradient posed significant
difficulties. Specifically, fractions containing the inhibitors would
also contain approximately 0.5 M sodium acetate, whose reaction
with o-phenylenediamine would give rise to benzimidazoles,
which could interfere with subsequent spectroscopic analyses of
the inhibitor-derived quinoxalines. Although this approach was
739
attempted, with subsequent sample clean-up by a combination of
solid phase extraction processes, it was not successful.
Instead, preparations of Cu2 + -oxidized RuBP were resolved
by anion-exchange chromatography, according to the method of
Kane et al. [12] (Protocol B), with minor changes to gradient
profile and flow rate, as appropriate for the larger MonoQ
(100 mm long × 10 mm diameter) column used in the present
study (see the Experimental section). Peaks were identified by
monitoring UV absorbance at 280 nm. This approach revealed a
minor peak at 58–60 min corresponding to RuBP and a major
peak at 76 min (Supplementary Figure S3, inset, available at
http://www.BiochemJ.org/bj/442/bj4420733add.htm). Fractions
collected in the vicinity of the major UV absorbing peak were
tested for the presence of Rubisco inhibitors and were consistently
found to contain two partially overlapping peaks of inhibition
(Supplementary Figure S3). The first inhibitor to emerge had
little (if any) UV absorbance, and was followed by a peak of
inhibition which coincided with the peak of UV absorbance. The
former peak of inhibition was likely to correspond to the first
peak of inhibition seen during the CarboPac PA1 fractionation of
the same material (Figures 6c and 6f) and the UV absorbing
inhibitor to the later eluting inhibitor identified by the same
method (Figures 6a and 6c). As they were collected, the fractions
corresponding to the leading edge of the earlier eluting inhibitor
(I1) and of the trailing edge of the later eluting inhibitor (I2),
denoted respectively by * and # (Supplementary Figure S3),
were combined with equal volumes of 0.1 M Hepps and 0.2 M ophenylenediamine, pH 8.0, followed by incubation in darkness for
60 min at room temperature, during which quinoxaline formation
could take place. Any phosphate-containing quinoxaline arising
in this way was purified by anion-exchange HPLC using a
MonoQ (100 mm long × 10 mm diameter) column and a simple
NaCl gradient, in the absence of any other solutes. Thus, for
each o-phenylenediamine-treated inhibitor (I1 or I2), a single
quinoxaline emerged, as a discrete and prominent peak with
absorbance maxima at both 238 nm and 319 nm (characteristic for
quinoxalines). These derivatives were concentrated, desalted by
passage through reverse-phase (C18 , end-capped) columns using
pure water as the mobile phase, and their identities determined by
NMR and MS.
The 1 H-NMR spectra from the quinoxalines of I1
and I2 were identical with each other, apart from their
concentration within the samples and differing amounts of
minor solvent impurities. Both contained peaks consistent
with the quinoxalines illustrated (Supplementary Figure S4
at http://www.BiochemJ.org/bj/442/bj4420733add.htm). Signals
from four aromatic protons were present as three signals between
δ 8.18 and 7.93. Remaining signals for two CH2 O(P) and one
CHOH group were evident between δ 5.62 and 4.20. A double
doublet at δ 5.62 with vicinal coupling to each of the protons on
C-11 was assigned to H-10. Remaining signals corresponded to
those adjacent to the phosphate groups and contained phosphorus
coupling of 6.5–7.0 Hz. Signals relating to the protons on C-9
appeared as a pair of double doublets centred at δ 5.374 and δ
5.328. Each signal contained couplings of 13 and 6.5 Hz relating
to geminal and phosphorus coupling respectively. Finally, the
multiplet appearing at δ 4.25 corresponded to hydrogens on C11 and actually comprises two closely located signals centred
at δ 4.286 and 4.236. These complex signals contained geminal
(11 Hz), vicinal and phosphorus (7 Hz) coupling for each of
the hydrogens. The multiplicity of each of the H-11 signals
was different, arising from different couplings to H-10 (7 Hz
or 4.5 Hz). The signal at δ 4.236 is therefore a pair of overlapping
triplets (J = 7 Hz), whereas the signal centred at δ 4.286 is a
double double doublet with 11, 7 and 4.5 Hz couplings.
c The Authors Journal compilation c 2012 Biochemical Society
740
P. J. Andralojc and others
Both samples gave an ion of m/z 401 in negative ion mode. In
addition, a smaller ion of m/z 379 was present in both samples
(Figure 7). Accurate mass analysis was carried out by direct
infusion on a Waters Q–TOF 1 MS. In the negative ion mode,
ions were observed at 400.9923 and 379.0111. m/z 400.9923
is consistent with empirical formula of C11 H12 N2 O9 P2 Na. This
would give a calculated mass of 400.9916 and thus the observed
ion represents an acceptable accuracy of 1.7 p.p.m. The formula
is consistent with an [M–2H + Na] − ion expected from the
quinoxaline derivative (as the sodium salt as shown). This is
in line with the ion obtained from RuBP (Mr 310), which
behaved in the same way and gave the similar [M–H] − ion
from the monosodium salt, at m/z 331. This phenomenon has
also been reported previously for ribuloselysine 3-phosphate
[31]. m/z 379.0111 is consistent with an empirical formula of
C11 H13 N2 O9 P2 . This would give a calculated monoisotopic mass
of 379.0096 and thus the observed ion represents a difference of
+ 4 p.p.m. The formula is consistent with [M–H] − that is expected
from the molecular ion of the quinoxaline derivative. There was
also one relevant fragment at 281.0291, which is consistent with
a [M–H] − ion arising from a compound of mass 282.03693
for which the suggested formula is C11 H11 N2 O5 P (calculated
mass = 282.0406, accuracy − 12.9 p.p.m). This represents a loss
of H3 PO4 from 380 or NaH2 PO4 from 402. MS/MS (tandem
MS) fragmentation is also highly consistent with the structure
and is given in Supplementary Figure S5 and Table S1 (at
http://www.BiochemJ.org/bj/442/bj4420733add.htm).
DISCUSSION
CA1Pase gene sequence
Peptide sequence information from P. vulgaris CA1Pase
combined with sequence data from a homologous Arabidopsis
gene, enabled the full-length homologous genes from bean,
tobacco and wheat to be cloned and sequenced (Figure 2).
The deduced protein sequence included a motif identical with the
PGM-active site sequence [LIVM]-x-R-H-G-[EQ]-x-x-[WN] at
the N-terminal end of the mature polypeptide. This sequence is
common to other enzymes involved in the transfer of phosphate
groups, such as FBPase (fructose bisphosphatase) and PFK
(phosphofructokinase). Structures for at least 36 polypeptides
with a similar active site sequence are available, including PGMs,
FBPases and PFKs. The CA1Pase sequence (Figure 2) appears to
contain two domains of roughly equal size, containing a number of
sequence homologies. The N-terminal PGM domain is followed
by a PFK-like domain. The occurrence of two such domains
in a single polypeptide is not without precedent: a bifunctional
mammalian enzyme, PFK/FBPase, possesses an N-terminal PFK
domain with an FBPase domain [32]. Although the overall amino
acid identity between the homologous sequences of Figure 2 is
51 % (228/451) significantly greater homology is apparent in the
N-terminal domain (66 %, 149/225) than the C-terminal domain
(35 %, 79/226) implying more stringent conservation of function
in the former domain. When current databases were scanned for
homologies of the wheat CA1Pase sequence, 12 plant accessions
were found with an amino acid sequence identity greater than
50 % (Table 1). Sequences from other monocots showed the
highest homology (76–80 % sequence identity), followed by dicot
sequences (60–65 % sequence identity), while those of a moss and
two unicellular green algae were more divergent. By contrast, the
length and sequence of the chloroplast transit peptides were not
highly conserved (Figure 2).
PGMs (EC 5.4.2.1) catalyse the transfer of phosphate groups
between carbon atoms 2 and 3 of phosphoglycerate, which
c The Authors Journal compilation c 2012 Biochemical Society
Table 1 Species, accession numbers and amino acid sequence identities
of gene sequences homologous with wheat CA1Pase
Species
Accession number
Amino acid identities
Sorghum bicolour
Zea mays
Oryza sativa Japonica (Os11g0150100)
Oryza sativa Indica group
Ricinus communis
Vitis vinifera
Populus trichocarpa
Arabidopsis thaliana (AT5G22620)
Micromonas sp. RCC299
Physcomitrella patens subsp. patens
Micromonas pusilla CCMP1545
Ostreococcus tauri
XM_002448951.1
ACR34191.1
NP_001065757.1
XP_002533602.1
XP_002533602.1
CBI17078.1
XP_002328174.1
NP_197654.1
XP_002503607.1
XP_001761673.1
EEH60229.1
CAL54665.1
357/446 (80 %)
188/234 (80 %)
356/446 (79 %)
320/421 (76 %)
293/448 (65 %)
295/449 (65 %)
290/450 (64 %)
267/439 (60 %)
136/235 (57 %)
263/466 (56 %)
128/233 (54 %)
120/222 (54 %)
is essential for the metabolism of glucose [33]. CA1Pase
was shown to have a diphosphoglycerate phosphatase activity
(Figure 3) indicating a functional resemblance to co-factordependent PGMs, although no bona fide PGM activity was
detected. Dephosphorylation of 2,3-DPG by CA1Pase yielded
2-PGA (2-phosphoglyceric acid). Similar to PGMs, the catalytic
mechanism of CA1Pase is likely to include the formation of a
phosphohistidine intermediate, possibly involving the histidine
residue of the highly conserved N-terminal sequence motif,
RHG (above). This is supported by our earlier observations
that CA1Pase purified from leaf extracts mediate a phosphate
exchange reaction, by which the phosphate group from nonradiolabelled CA1P is transferred to [14 C]CA, consistent with
the formation of a phosphoenzyme intermediate [17].
The expressed protein is a CA1Pase
The properties of the protein encoded by the cloned CA1Pase
gene from wheat are broadly consistent with those of the enzyme
purified from French bean and tobacco [21,26–29]. The V max for
CA1P, 4.2 μmol/min per mg of protein, was very similar to that
of the counterpart from P. vulgaris of 6–7 μmol/min per mg
of protein [26,28]. The higher values reported for P. vulgaris
may reflect interspecies differences, but may also be due to the
reported inclusion of either DTT plus KCl [26] or of DTT plus
FBP [28] in the respective assay medium, both of which have
been shown to stimulate catalytic activity. However, none of
these effectors were present in the determinations of Figure 3.
Comparison of K m values (Figure 3) indicates higher affinity
for bisphosphate analogues of CA1P, than for CA1P itself, the
values being 2–3-fold lower than for CA1P. The K m for CA1P
reported in the present paper is very similar to the 12.6 +
− 2.0 μM
reported for the tobacco enzyme [30], but is very much lower
than the 433 +
− 26 μM for P. vulgaris [26]. This difference may
reflect the different concentrations of CA1P known to occur in
the two species: high in P. vulgaris, moderate in N. tabacum and
low in wheat [7]. Our determinations of the kinetic constants of
CA1Pase (Figure 3) were conducted at the pH optimum of the
enzyme (pH 7.0) although investigations into the effect of other
metabolites on activity (Figure 4B) were performed at pH 8.0, as
this was more likely to reflect the stromal pH when many of the
chosen effectors would be present (see below).
The effect of phosphate-containing metabolites on the rate
of CA1P breakdown catalysed by CA1Pase are in agreement
with earlier studies, with the notable exception of 2,3-DPG
CA1P phosphatase and Rubisco regulation
(Figure 3) which had been reported to greatly stimulate the
dephosphorylation of CA1P [28]. However, since the phosphatase
activity in this previous study [28] relied on the detection of Pi ,
interpretation of the effects of other phosphorylated compounds
on the dephosphorylation of CA1P was ambiguous, since there
was no way of distinguishing between the loss of phosphate from
CA1P and the loss of phosphate from any other organic phosphate.
The fact remains that many phosphorylated metabolites which
are not substrates themselves do stimulate CA1P breakdown
by CA1Pase [26,28,30] (Figure 4B). The likelihood that
CA1Pase is composed of two distinct functional domains
(discussed above) suggests that CA1Pase activity resides on
one domain, while the other may interact with a variety of
phosphorylated effectors, bringing about changes in the rate
of CA1P dephosphorylation.
There are eight conserved and 13 species-specific cysteine
residues in the aligned primary sequences of Figure 2. Differing
reports exist regarding the extent of activation of the enzyme
by the sulfhydryl-containing reagent, DTT. A slight activating
effect of DTT on CA1Pase activity was reported in P. vulgaris
[26], although in the presence of KCl it was mildly inhibitory [26].
Conversely, a large activating effect of DTT on the activity in
N. tabacum has been demonstrated [30]. More recent reports
suggest a very large stimulation of the enzyme from P. vulgaris
by GSH [34]. The differences in the distribution of cysteine
residues are likely to account for some of these differing
responses. The catalytic activity of any CA1Pase preparation
in the absence of exogenous oxidant or reductant is likely to reflect
the immediate history of the preparation (e.g. the prevalence
of oxidizing or reducing conditions during its isolation). The
change in activity between the fully oxidized and reduced
states, however, indicates the extent to which the activity of the
enzyme can be redox modulated. In preliminary experiments,
we found that 10 mM DTT stimulated CA1Pase activity at least
to the same extent as 10 mM GSH, and so utilized DTT as
reductant thereafter. The susceptibility of each of the CA1Pase
preparations to redox mediated changes in activity were very
pH-sensitive, there being very little redox modulation at pH 6,
an intermediate response at pH 7 and very considerable redox
modulation at pH 8 (Figure 4A), consistent with pH-dependent
conformational changes determining the accessibility of redoxsensitive cysteines. The pH of the chloroplast stroma changes
between pH 7 in the dark to pH 8 in the light [2] and the
redox potential of the stroma becomes more reducing during
periods of illumination [2]. The pronounced redox sensitivity at
pH 8 suggests that redox-mediated changes in CA1Pase activity
during the day have a role in vivo, for example during transient
adaptations to a fluctuating light environment. However, the
apparent lack of an effective redox switch at pH 7 would imply that
CA1Pase activity would be relatively high in the dark, when CA1P
production is likely to be at its height, reducing the availability
of CA1P for binding to Rubisco and/or leading to a futile
cycle.
Activity of CA1Pase towards other Rubisco inhibitors
The two Rubisco inhibitors derived from RuBP under oxidizing
conditions have retention times during HPLC consistent with
bisphosphates. They also contaminate commercial preparations
of RuBP (Figure 5 and Supplementary Figure S2). Based on
the A280 and either the phosphate content or the inhibitory
activity of the HPLC-resolved peak of the later eluting RuBP
derivative (Figure 6a), a molar absorption coefficient in excess of
104 M − 1 cm − 1 was calculated. For comparison, we separately
741
determined molar absorption coefficients (280 nm, 25 ◦ C, in
water) of 46 and 7 M − 1 · cm − 1 for RuBP and CABP respectively,
and of 1.5 × 104 M − 1 · cm − 1 for 3,4-dihydroxy-3-cyclobutene1,2-dione whose vicinal di-keto moiety is flanked by hydroxylbearing carbons, as in PDBP. Thus the high absorbance at
280 nm of the later eluting inhibitory peak is consistent with
compounds containing vicinal keto groups flanked on either
side by hydroxylated carbons. The absence of absorbance at
280 nm of the earlier eluting inhibitory peak would be expected
for a hydrated form of PDBP, resulting from an equilibrium of
the form >C = O + H2 O↔>C(OH)2 . The hydration of ketones
is a widely recognized phenomenon (reviewed in [35]). Such
hydration would be accompanied by the disappearance of the
vicinal di-keto moiety and thus a fall in absorbance at 280 nm.
Although hydration of ketones would be diminished by the
proximity of large substituents due to steric hindrance, it would be
favoured by the proximity of electron-withdrawing groups such
as a neighbouring carbonyl moiety, which may favour hydration
by destabilizing the carbonyl group on C-2. Both PDBP and
its hydrated counterpart would be expected to form identical
2, 3-substituted quinoxalines, and this has been demonstrated
for the two partially overlapping inhibitory peaks shown in
Supplementary Figure S3. Such a simple relationship between
the two inhibitors would also provide an explanation for the
observed interconversion between the two chromatographically
distinct inhibitors demonstrated in Figure 6. The ability of the
CarboPac PA1 column to distinguish between these two forms
of PDBP would be expected for a column which had been
developed to differentiate between closely related sugars and their
phosphates. We conclude that the maintenance of Rubisco activity
promoted by the presence of CA1Pase (Figure 5) was due to the
dephosphorylation of contaminating PDBP (or its corresponding
hydrate) by CA1Pase.
Neither RuBP nor XuBP was found to be a substrate for
CA1Pase. Since PDBP is structurally very similar to these sugar
bisphosphates, it is surprising that it should be a substrate for
CA1Pase (Figure 1). However, hydration of the carbonyl group
of C-2 may yield a structure that is dephosphorylated more
readily by CA1Pase. Since the two inhibitors are interconvertible,
the dephosphorylation of one of the inhibitors would result in the
gradual disappearance of the other inhibitor without the need for
direct interaction between both forms of PDBP and CA1Pase.
A putative coding sequence for CA1Pase has been found in
plant species (Figure 2 and Table 1) irrespective of the occurrence
of photosynthetically significant amounts of CA1P. For example,
wheat and Arabidopsis have very little (if any) CA1P, while French
bean, tobacco and rice contain considerable amounts of CA1P
[7,8,36] and yet each of these species possess a putative gene
for CA1Pase (Figure 2 and Table 1). Indeed, CA1Pase activity is
significant [28] in plant species with little CA1P (including wheat
and Arabidopsis), albeit less than that detected in French bean or
potato [28]. This indicates a role for this enzyme in processes
distinct from CA1P removal. The observation that CA1Pase
maintains the activity of Rubisco by removing contaminating
inhibitors, likely to be derived from RuBP in the presence of
oxygen [12,13], indicates an alternative role. It remains to be
seen whether such RuBP-derived inhibitors can be produced nonenzymically in vivo, but their occurrence as by-products of side
reactions catalysed by Rubisco in vitro has been documented
[12–14] and evidence for its occurrence in illuminated wheat
leaves in vivo has been presented [37]. The observation that
CA1Pase has a higher affinity for the bisphosphate analogues
of CA1P than for CA1P itself (Figure 3) and that CA1Pase purified
from P. vulgaris and N. tabacum dephosphorylate CABP faster
than CA1P [26–28] supports this hypothesis. The observation that
c The Authors Journal compilation c 2012 Biochemical Society
742
P. J. Andralojc and others
the K m for CABP is in the low micromolar range suggests that,
if the K m for an analogous inhibitory derivative of RuBP is
similarly low, then the accumulation of such compounds to the
extent that they would account for a significant proportion of
the available Rubisco catalytic sites would be unlikely.
Our results are consistent with an additional role for CA1Pase in
the removal of PDBP. We envisage that this role for CA1Pase
would be accomplished in conjunction with Rubisco activase,
which is responsible for the prerequisite release of inhibitors from
the active site of Rubisco. In this way, the occurrence of Rubisco
fallover could be controlled in vivo.
AUTHOR CONTRIBUTION
Jane Loveland purified CA1Pase from P. vulgaris and generated CA1Pase peptides. Phil
Jackson and Antony Willis sequenced peptide fragments from CA1Pase. Pippa Madgwick,
Yong Tao and Steven Gutteridge identified, expressed and confirmed the function of
the CA1Pase gene from wheat. Pippa Madgwick was responsible for all other genetic
manipulation and sequencing procedures. Paul John Andralojc was responsible for the
illustrated experiments. Mike Beale and Jane Ward were responsible for the NMR and MS
analyses. Alfred Keys provided expertise on substrate preparation and PDBP. Paul John
Andralojc wrote the paper. Martin Parry supervised the study.
ACKNOWLEDGEMENTS
We thank Brian G. Forde (Lancaster University) for his assistance in identifying a putative
CA1Pase gene; Heather Kane (Australian National University) for providing information
that enabled the synthesis of XuBP; and John Baker (Rothamsted Research) for ESI–MS
data collection.
FUNDING
Rothamsted Research receives grant-aided support from the Biotechnological and
Biological Sciences Research Council of the U.K.
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Biochem. J. (2012) 442, 733–742 (Printed in Great Britain)
doi:10.1042/BJ20111443
SUPPLEMENTARY ONLINE DATA
2-Carboxy-D-arabinitol 1-phosphate (CA1P) phosphatase: evidence for
a wider role in plant Rubisco regulation
Paul John ANDRALOJC*1 , Pippa J. MADGWICK*, Yong TAO†2 , Alfred KEYS*, Jane L. WARD*, Michael H. BEALE*,
Jane E. LOVELAND*3 , Phil J. JACKSON‡, Antony C. WILLIS§, Steven GUTTERIDGE† and Martin A.J. PARRY*1
*Department of Plant Sciences, Rothamsted Research, Harpenden AL5 2JQ, U.K., †DuPont Stine-Haskell Research Center, Newark, DE 19714, U.S.A., ‡Chemical and Process
Engineering, University of Sheffield, Sheffield S3 7RD, U.K., and §MRC Immunochemistry Unit, University of Oxford, Oxford OX1 3QU, U.K.
Figure S1 Time and concentration dependence of CA1P dephosphorylation
by the expression product of the wheat homologue of At5g22620 (CA1Pase)
The inset illustrates the purity and molecular mass (MW) in kDa of this protein as revealed by
SDS/PAGE using a 4–20 % gradient gel (Pierce) after staining with Coomassie Brilliant Blue
(1 μg of the CA1Pase was loaded).
Figure S2
Removal of inhibitory contaminants in RuBP by CA1Pase
An aliquot (0.5 ml) of a 4 mM solution of the commercial RuBP preparation was pre-incubated
for 30 min in a buffer containing 25 mM Bicine (pH 8.0), 1 mg BSA/ml, 0.5 mM EDTA and
5 mM DTT in the presence (B) or absence (A) of 10 μg of DTT-activated CA1Pase, followed by
protein removal by ultrafiltration (using Centricon YM-10 centrifugal concentrators) and HPLC
analysis/fractionation. Fractions collected at 0.5 min intervals from the emergence of RuBP
onwards were tested for the presence of Rubisco inhibitors by measuring their effect on Rubisco
carboxylase activity, as shown by the overlaid histograms, which show the mean and S.D. for
duplicate determinations.
1
2
3
Correspondence may be addressed to either of these authors (email [email protected] or [email protected]).
Present address: Institute of Microbiology, Chinese Academy of Sciences, Beijing 100101, People’s Republic of China.
Present address: Wellcome Trust Sanger Institute, Hinxton CB10 1HH, U.K.
c The Authors Journal compilation c 2012 Biochemical Society
P. J. Andralojc and others
Figure S3 Partial resolution of two RuBP-derived Rubisco inhibitors by
anion exchange HPLC prior to quinoxaline formation
Cu2 + /O2 -oxidized RuBP (1 ml) was resolved using a MonoQ (10/10) column with NaCl
gradient elution and fraction collection at 0.5 min intervals. Then 2.5 μl aliquots of the fractions
indicated were taken immediately (for assay of Rubisco inhibitors) and an equal volume of the
o -phenylenediamine reagent promptly added to the remainder. Inhibitory fractions marked ‘∗’
(I1) or ‘#’ (I2) were combined and subsequently analysed by NMR and MS. The inset shows the
whole elution profile.
c The Authors Journal compilation c 2012 Biochemical Society
CA1P phosphatase and Rubisco regulation
Figure S4
1
H-NMR spectra of the quinoxalines of I1 and I2
These data can be summarized as: 1 H-NMR (H2 O, 600 MHz): δ 8.18 (1H, m, H-5/H-8), 8.14 (1H, m, H-5/H-8), 7.93 (m, 2H, H-6 and H-7), 5.59 (1H, dd, 4.5 Hz and 7 Hz, H-10), 5.37 (1H, dd, 6.5 Hz
and 13 Hz, H-9a ), 5.33 (1H, dd, 6.5 Hz and 13 Hz, H-9b ), 4.29 (1H, ddd, 4.5 Hz, 7 Hz and 11 Hz, H-11a ), 4.24 (1H, dt, 7 Hz and 11 Hz, H-11b ).
c The Authors Journal compilation c 2012 Biochemical Society
P. J. Andralojc and others
Figure S5
MS/MS fragmentation analysis
MS/MS was carried out on a Bruker Ion-trap and the results were identical for I1 and I2, providing
further confirmation that both fractions contained the same metabolite. (A) Fragmentation of
the sodium salt, m /z 401, showed a complicated fragmentation pattern. The MS2 spectrum
contained major product ions at m /z 383 (100 %) and 371 (97 %) and further fragments at
281 (30 %), 263 (2 %), 251 (10 %) and 199 (67 %). Further MS3 fragmentation of 383 and
371 revealed that there were two different pathways of fragmentation of 401, both giving rise
to 199. m /z 383 also showed losses of 102 and 120 to give rise to the m /z 281 and 263 ions
respectively. M /z 371, in addition to the breakdown to 199 directly, also showed the loss of 102
and 120 to give ions at m /z 269 and 251. The losses represent H2 O (18), CH2 O (30), NaPO3
(102), NaH2 PO4 (120). Losses of 18,102 and 120 are obvious, but the loss of 30 indicates a
‘free’ terminal CH2 OH. From the phosphorus coupling observed in NMR, we know that both
phosphate groups are attached to CH2 O- and not to the -CH–O-. It can therefore be postulated
that the loss of 30 from the molecular ion of the sodium salt results from a concerted process
in the gas phase, whereby the phosphate group shifts to the secondary hydroxyl position and
the C-C bond is cleaved (B). The structure of m /z 199, however, was not clear. This ion comes
from MS/MS of m /z 401 directly and can also be obtained via MS/MS of both m /z 383 and
371. We have not been able to determine any pathway, via MS/MS, to m /z 199 from m /z 281,
263, 269 or 251. Further MS/MS of m /z 400.99 on the Q–TOF was used to determine possible
formulae for the key fragments. From this we have determined that the m /z 199 is due to
sodium diphosphate. Further related fragments, 181, 97 and 79 result from this ion. The loss of
diphosphate only occurs from the heavier ions 401, 383 and 371. The complete fragmentation
tree derived from Ion-trap MS/MS and Q–TOF MS/MS can thus be deduced (C). The smaller
[M –H] − ion in the original ESI (electrospray ionization)–MS spectrum (m /z 379) gave a much
simpler fragmentation pattern and showed a product ion at m /z 281 which is a loss of 98 atomic
mass units, consistent with loss of H3 PO4 . No further MS/MS of m /z 281 was observed.
c The Authors Journal compilation c 2012 Biochemical Society
CA1P phosphatase and Rubisco regulation
Table S1
Mass and structural formulae of ion fragments identified by MS/MS
Measured mass (Da)
Empirical formula
Calculated mass (Da)
Accuracy (p.p.m.)
Structure
O
OP
O
OH
N
N
O
OH
382.9985
C11 H10 O8 N2 P2 Na
382.9810
P
O
ONa
45
O
-
O
P
O
OH
N
N
ONa
O
P
OH
370.9987
C10 H10 O8 N2 P2 Na
370.9810
47
O
O
OP
O
OH
N
HO
299.0489
C11 H12 O6 N2 P
299.0433
N
OH
19
O
OP
O
OH
N
281.0306
C11 H10 O5 N2 P
281.0327
7 ppm
N
HO
O-
O
O
P
P
HO
198.9225
180.9150
96.9688
78.9489
NaH2 P2 O7
P2 O6 Na
H2 PO4
PO3
198.9173
180.9068
96.9691
78.9585
26
45
3
121
O
OH ONa
Received 5 August 2011/21 November 2011; accepted 2 December 2011
Published as BJ Immediate Publication 2 December 2011, doi:10.1042/BJ20111443
c The Authors Journal compilation c 2012 Biochemical Society