Download Purification and Characterization of

Plant Physiol. (1998) 117: 321–329
Purification and Characterization of Phosphoribulokinase
from the Marine Chromophytic Alga Heterosigma carterae1
Tara Hariharan, Paula J. Johnson, and Rose Ann Cattolico2*
Department of Botany (T.H., P.J.J., R.A.C.), and School of Oceanography (R.A.C.), University of Washington,
Seattle, Washington 98195
In this study we characterized phosphoribulokinase (PRK, EC
2.7.1.19) from the eukaryotic marine chromophyte Heterosigma
carterae. Serial column chromatography resulted in approximately
300-fold purification of the enzyme. A polypeptide of 53 kD was
identified as PRK by sequencing the amino terminus of the protein.
This protein represents one of the largest composite monomers
identified to date for any PRK. The native holoenzyme demonstrated
by flow performance liquid chromatography a molecular mass of
214 6 12.6 kD, suggesting a tetrameric structure for this catalyst.
Because H. carterae PRK activity was insensitive to NADH but was
stimulated by dithiothreitol, it appears that the enzyme may require
a thioredoxin/ferredoxin rather than a metabolite mode of regulation. Kinetic analysis of this enzyme demonstrated Michaelis constant
values of ribulose-5-phosphate (226 mM) and ATP (208 mM), respectively. In summary, H. carterae PRK is unique with respect to
holoenzyme structure and function, and thus may represent an alternative evolutionary pathway in Calvin-cycle kinase development.
Three mechanisms have been described by which CO2
can be autotrophically processed. The reductive citric acid
cycle and acetyl CoA condensation reaction are found exclusively in the green, methanogenic, and acetogenic bacteria (Hemming and Blotevogel, 1985; Fuchs, 1986; Schäfer
et al., 1989). In contrast, the Calvin cycle is used by diverse
organisms, including bacteria and eukaryotes, for CO2 processing (McFadden and Tabita, 1974).
Carbon isotope fractionation indicates that the Calvin
cycle has been used to process CO2 for more than 3.5 billion
years (Raven, 1997). The origin of this cycle is not well
understood. Two enzymes, Rubisco and PRK, are unique to
Calvin-cycle function and they probably provided the evolutionary “breakthrough” with respect to the biogenesis of
this metabolic pathway, because the remaining Calvincycle enzymes have additional (i.e. non-Calvin cycle) metabolic responsibilities.
PRK catalyzes the reaction Ru5P 1 ATP 3 ribulose-1,5bisphosphate 1 ADP. Because this reaction is essentially
irreversible, the enzyme serves a critical role in regulating
the flow of sugars through the CO2-fixation cycle. Some of
the first catalysts to occur in primitive cells were similar to
1
Supported by National Science Foundation grant no. MCB9305923.
2
All authors contributed equally to this study.
* Corresponding author; e-mail [email protected]; fax
1–206 – 685–1728.
extant kinases (Loomis, 1988). These enzymes phosphorylated a wide range of molecules with ATP. Thus, unlike
Rubisco, for which no other enzymatic analog exists
(McFadden and Tabita, 1974), one may hypothesize that
PRK was probably a “retrofitted” kinase. Any enzyme that
was capable of moving a phosphate from ATP to an acceptor molecule can serve as a candidate for Calvin-cycle
specialization.
Kinases tend to be monomers. However, PRK does not fit
this profile, perhaps as a result of Calvin-cycle isolation
(Loomis, 1988). The structure of PRK varies extensively
among taxa. Terrestrial plants and algal representatives of
the phylum Chlorophyta (chl a- and b-containing plants)
have the simplest PRK design. In these taxa (Table I), 80- to
90-kD holoenzymes are homodimeric except in Selenastrum
minutum, which is heterodimeric (Lin and Turpin, 1992).
Spinach PRK exists in the chloroplast stroma in a multienzyme complex (Gontero et al., 1988). All enzymes of this
complex are committed to Calvin-cycle function. A similar
but smaller multienzyme complex has been described for
pea (Sainis and Harris, 1986).
The structural identity of PRK among prokaryotes is
taxon dependent (for review, see Table I). Cyanobacterial
PRK holoenzyme composition is quite variable. For example, the PRK of Anabaena cylindrica is slightly smaller (72
kD) than that found in terrestrial plants. Both 43- and
26-kD composite proteins were observed. It is not known
whether the smaller protein represents a subunit of the
holoenzyme or a tightly associated contaminating polypeptide. Synechococcus spp. and Chlorogleopsis fritschii PRK both
display monomeric subunits of 40 kD. The Synechococcus
spp. enzyme, however, is tetrameric in structure (178 kD),
whereas the C. fritschii enzyme is hexameric (230 kD).
Cyanobacterial PRK holoenzyme structural differences are
not seen in proteobacterial PRK enzymes, which exhibit
relatively uniform molecular masses. These bacteria contain a holoenzyme of 250 kD that is composed of six to
eight subunits, each with a molecular mass of approximately 35 kD.
Although the data are not entirely representative of all
cells that use the Calvin cycle for CO2 fixation, it appears
that PRK regulation occurs by two different mechanisms.
These differences in PRK regulation may reflect separate
Abbreviations: chl, chlorophyll; DE-52, diethylaminoethyl cellulose; PRK, phosphoribulokinase; Rib5P, ribose-5-phosphate;
Ru5P, ribulose-5-phosphate.
321
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322
Hariharan et al.
Plant Physiol. Vol. 117, 1998
Table I. Summary of PRK structure and regulatory control requirements among divergent taxa
Organism
Holoenzyme
Monomer
Subunit
D
Chlorophyta
Arabidopsis
Ice plant
Spinach
Wheat
Bryopsis maxima
Chlamydomonas reinhardtii
Chlorella sorokiniana
Scenedesmus obliquus
S. minutum
NDa
ND
90,000
ND
90,000
Proteobacteria
Alcaligenes eutrophus
Agmenellum quadruplicatum
Chromatium sp.
Hydrogenomonas eutropha H16
Nitrobacter winogradskyi
Rhodobacter sphaeroides
Rhodopseudomonas acidophila
Rhodopseudomonas capsulata
Rhodopseudomonas spaeroides
Thiobpacillus neapolitanus
Xanthobacter flavus H4-14
a
ND, Not determined.
encoded.
b
NADH
Ref.
n
ND
ND
2
ND
2
ND
ND
2
2
ND
ND
1c
1
ND
1
ND
1
ND
ND
ND
ND
ND
ND
ND
2d
ND
ND
Horsnell and Raines (1991)
Michaelowski et al. (1992)
Porter et al. (1986)
Raines et al. (1989)
Satoh et al. (1985)
Roesler and Oregen (1990)
Tabita (1980)
Lazaro et al. (1986)
Lin and Turpin (1992)
2
1
ND
Serra et al (1989)
230,000
ND
178,000
43,000
26,000
40,000
39,000
42,000
6
ND
4
1
1
1
2
ND
ND
Marsden and Codd (1984)
Su and Bogorad (1991)
Wadano et al. (1995)
214,000
ND
53,000
45,000b
4
ND
1
ND
2
ND
This work
K. Weyrauch (1996) Genbank accession no.
Y08610
256,000
33,319e
33,164f
ND
ND
ND
ND
33,000
32,000
36,000
33,000
ND
33,409
8
ND
ND
Siebert et al. (1981); Kossmann et al. (1989)
ND
ND
ND
ND
ND
8
6
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
ND
1
2
ND
1
1
1
ND
1
1
ND
Tabita (1980)
Hart and Gibson (1971)
Abdelal and Schlegel (1974)
Kiesow et al. (1977)
Hallenbeck and Kaplan (1987)
Rippel and Bowien (1984)
Tabita (1980)
Hallenbeck and Kaplan (1987)
Tabita (1980)
Kossman et al. (1989); Meijer et al. (1990)
72,000
C. fritschii
Synechocystis sp. PCC 6803
Synechococcus sp. PCC 7942
Chromophyta
Heterosigma carterae
Odentella sp.
DTT
38,400b
44,000b
44,000
39,200b
41,000
39,000
ND
42,000
41,000
42,000
83,000
83,000
Cyanobacteria
A. cylindrica
Activation
ND
240,000
237,000
ND
248,000
220,000
ND
ND
ND
Inferred from DNA sequence data.
evolutionary origins or early evolutionary divergence of
PRK. Many but not all bacterial PRK enzymes are strongly
activated by NADH (Table I) (Gibson and Tabita, 1987).
PRK activation via this type of allosteric modulation may
be quite effective (Kiesow et al., 1977) because energylinked NADH-generating reactions (e.g. via nitrate oxidation) are functional in both photosynthetic and chemoautotrophic bacteria. In contrast, chlorophytes (chl a- and
b-containing plants) and cyanobacteria use thioredoxin in
the modulation of Calvin-cycle enzymes, including PRK
(Holmgren, 1985; Hartman et al., 1990). In the light, electrons from chl are transferred to Fd and then to thioredoxin. The reduced thioredoxin activates PRK by affecting
the redox-active S-S bridge of the enzyme. Milanez et al.
(1991) has shown that regulation of PRK activity involves
Cys residues 16 and 55 in the protein. Sequence analysis
has shown that equivalent Cys residues are not found in
allosterically regulated proteobacterial PRK.
The Chromophyta (chl a- and c-containing plants) represent a large assemblage of organisms that rank as the
c
1, Present.
d
2, Absent.
e
Chromosome encoded.
f
Plasmid
primary producers in many aquatic ecosystems. Chromophytes have a significant effect on the global carbon budget. Approximately one-third of the total carbon fixed
worldwide is processed by these organisms (Raven, 1997).
To our knowledge, there has been no analysis of PRK
structure and function in a marine eukaryote. Here we
present information on the isolation and characterization of
PRK from the toxic, unicellular marine alga H. carterae.
MATERIALS AND METHODS
All chemicals, enzymes, and column supports were obtained from Sigma except for MgCl2 and KCl (J.T. Baker),
Tris base (GIBCO-BRL), Sephadex G-50 and DE-52 cellulose (Whatman), and a Superose-6 flow performance liquid
chromatography column (Pharmacia). Antibodies to Alcaligenes eutrophus PRK were kindly provided by B. Bowien
(Universität Göttingen, Germany).
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Phosphoribulokinase in Marine Chromophytic Alga
Algal Culture
Heterosigma carterae (Taylor, 1992), isolate Carter, was
grown in an artificial seawater medium (McIntosh and
Cattolico, 1978) at 20°C on a 12-h light/12-h dark diel cycle.
Cultures of 1 L were maintained in 2.8-L Fernbach flasks
with continuous shaking at 60 rpm. Cultures were illuminated with cool-white fluorescent lamps at an intensity of
20 mE m22 s21. Cells were counted using a ZBI counter
(Coulter, Hialeah, FL) with a 100-mm aperture.
Enzyme Purification
Unless otherwise indicated, all of the following procedures were done at 5°C. Cells were harvested when cultures had a density of approximately 105 cells mL21 by
centrifugation at 1,110g for 10 min. The cells were resuspended to a final concentration of 5 3 107 cells mL21 in 50
mm Hepes buffer (pH 7.8) that contained 10 mm MgCl2,
0.1% b-mercaptoethanol, and 1 mm PMSF as a protease
inhibitor. This mixture was stored at 270°C for later use.
An atmosphere of N2 was maintained over the homogenate
at all phases of cellular disruption and centrifugation. Approximately 4 3 109 stored cells (40 L of cells at 105 cells
mL21) were thawed on ice, diluted to a final concentration
of 2 3 107 cells mL21 using an equilibration buffer (50 mm
Tris [pH 7.6], 0.5 mm EDTA, 100 mm KCl) that contained
0.1% b-mercaptoethanol, and further disrupted by a single
passage through a precooled French pressure cell at
8,000 p.s.i. The broken cells were centrifuged at 10,900g for
15 min. Retrieved supernatant was then centrifuged at
100,000g for 40 min. A degassed solution that contained
saturated (NH4)2SO4 (pH 7.6) and 0.1% b-mercaptoethanol
was slowly added to the 100,000g supernatant to attain 35%
final concentration. This mixture was stirred for 30 min.
The resulting precipitate was removed by centrifugation at
8,700g for 30 min. Saturated (NH4)2SO4 was added to the
supernatant to achieve a final concentration of 50%, after
which solid (NH4)2SO4 was added to 80% final concentration. The solution was subject to continuous stirring for 30
min. The precipitate, retrieved by centrifuging at 8,700g for
30 min, was resuspended in 10 mL of equilibration buffer
containing 0.1% b-mercaptoethanol. The suspension was
flushed with N2. The 35 to 80% (NH4)2SO4 fraction was
desalted by passage through a Sephadex G-50 column (approximately 60 mL) that had been swollen in water and
washed with the same buffer. Fractions containing the
enzyme were further desalted by dialysis against the same
buffer for a minimum of 6 h. The dialyzed solution was
then subject to chromatographic separation (at room temperature) using DE-52 microcellulose anion-exchange resin
(Whatman) that had been swollen overnight in water,
washed with equilibration buffer, and then activated for 60
min by adding 500 mm KCl to the equilibration buffer.
The column was packed to a bed volume of 25 mL in
equilibration buffer containing 1 mm DTT using a pressure
pump set at 100 mL h21. The sample was loaded at 50 mL
h21 and the column was washed with six bed volumes of
equilibration buffer. Protein was eluted at 50 mL h21 using
a linear salt gradient of 100 to 400 mm KCl in equilibration
323
buffer containing 1 mm DTT. Fractions showing enzyme
activity were pooled and concentrated to one-tenth the
volume by high-pressure ultrafiltration using an Amicon
(Beverly, MA) model 12 stirred cell equipped with a PM30
Diaflo membrane at a maximum pressure (60 p.s.i.) of N2.
The sample was activated by adding 10 mm DTT, flushed
with N2, and stored on ice for 2 h.
Agarose-Reactive Green 19 that had been swollen in
water for several hours was equilibrated with 10 mm
Bicine-KOH buffer (pH 8.0) and then reequilibrated with 10
mm Bicine-KOH (pH 6.8) buffer that contained 10 mm
MgCl2. The pH of the PRK sample that had been activated
was adjusted to 6.8 with Bicine-KOH (pH 5.0). The sample
was then added to this affinity slurry and maintained at
4°C for 18 h to allow complete binding of the enzyme. The
Reactive Green 19 slurry containing bound enzyme was
brought to room temperature and packed into a column
that was washed with 10 bed volumes of 20 mm Tris-HCl
(pH 7.6), 10 mm DTT. Enzyme was eluted with 10 mm ATP
in the same buffer. Retrieved enzyme was further concentrated by ultrafiltration as described above. The recovered
protein was stored under N2 at 270°C after the addition of
10% glycerol, 1 mm leupeptin, and 10 mm DTT. This fraction was used for enzyme characterization and electrophoretic analyses.
Active fractions from the affinity column were chromatographed on a Superose-6 gel-filtration column that was
equilibrated with buffer composed of 50 mm Tris (pH 7.6),
0.5 mm EDTA, 100 mm KCl, 10 mm DTT. For molecular
mass determination of the native enzyme size, the
Superose-6 column was calibrated with catalase (232 kD),
aldolase (158 kD), BSA (68 kD), and Cyt c (12.5 kD). The
following equation was used to calculate the elution constant (Kav) of both PRK and the protein standards: Kav 5
(Ve 2 Vo)/(Vt 2 Vo), where Ve is the elution volume of
sample, Vo is the void volume, and (Vt 2 Vo) is the volume
of gel-forming substance (Pharmacia Biotech, 1993). Protein profiles were spectrophotometrically monitored during chromatography at 280 nm. Protein was quantified
using a Bio-Rad assay kit with BSA as a standard.
Protein Electrophoresis and Sequencing
Fractions showing PRK activity from the DE-52 column
and the Reactive Green 19 column were pooled, concentrated, and resolved on 12% SDS-PAGE gels (Laemmli,
1970). Gels were either stained with Coomassie blue R-250
or silver stained. PRK subunit size was determined by
comparison with known standards.
Proteins (10 mg) in the PRK-containing pool from a postReactive Green 19 column were separated on SDS-PAGE,
transferred to a PVDF membrane, and stained with Coomassie blue R-250. The major band was sequenced by
Edman degradation at the amino terminus (Protein Sequencing Facility, University of Washington, Seattle).
Enzyme Assays
Two coupled assays were used to assess PRK activity
during the course of this study. In the radioactive proce-
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324
Hariharan et al.
dure, the conversion of Ru5P to a three-carbon sugar was
assessed by monitoring the incorporation of 14C into an
acid-precipitable product (Paulsen and Lane, 1966). The
reaction mixture consisted of 0.5 mg of PRK extract, 50 mg
of spinach Rubisco, 50 mm Tris (pH 8.0), 20 mm DTT, 20%
glycerol, 30 mm NaHCO3, and 10 mm MgCl2 in 50 mL,
which was incubated on ice for 30 min. The volume of the
reaction mixture was increased to 200 mL, and the mixture
was adjusted to a final concentration of 100 mm Tris (pH
8.0), 5 mm ATP, 10 mm MgCl2, 20% glycerol, and 20 mm
DTT. This new solution was incubated on ice for 15 min,
after which 1 mL (0.06 Ci/mol) of NaH14CO3 was added.
The reaction was initiated with 2 mm Ru5P and incubated
for 10 min at 30°C. The assay mixture was then added to
200 mL of 2 n HCl that was contained in a scintillation vial,
brought to dryness by heating in a 95°C water bath, dissolved in 200 mL of distilled water, and the product was
counted in 3 mL of scintillation fluid (Ecolume, ICN) using
a scintillation counter (Beckman). Parameters of pH and
temperature were optimized for this assay as well as for the
spectrophotometric assay.
In the spectrophotometric assay, the activity of PRK was
analyzed via a coupled reaction (Kagawa, 1982). Use of
ATP during the phosphorylation of Ru5P by PRK was
coupled to the conversion of PEP to lactate. The oxidation
of NADH in this scheme was monitored at 340 nm. The
reaction mixture (1.0 mL) contained 100 mm Tris (pH 8.0),
3 mm MgCl2, 10 mm DTT, 2 mm PEP, 2 mm ATP, 4 mm
Rib5P, phosphoriboisomerase (2 units), lactic dehydrogenase (8 units), pyruvate kinase (13 units), and 0.1 mm
NADH. To eliminate effects of contaminating ADP, the
assay mixture minus PRK and Rib5P was incubated at 25°C
for 1 min. Background absorbance was observed by incubating the reaction mixture plus 5 mg of PRK enzyme. The
reaction was initiated by the addition of Rib5P. Change in
absorbance was monitored using a UV-160 spectrophotometer (Shimadzu Scientific, Columbia, MD) at 340 nm set on
kinetic mode. One unit of activity was defined as the
amount of enzyme that catalyzed the oxidation of 1 mmol
of NADH per minute (or production of ADP). To determine the Km of substrate, commercially available Ru5P was
added to the reaction mixture (instead of converting Rib5P
to Ru5P with phosphoriboisomerase, as is routinely done
for activity assays).
Plant Physiol. Vol. 117, 1998
RESULTS
Enzyme Isolation and Physical Characteristics
Although initial PRK isolations were done with isolated
chloroplasts, subsequent work demonstrated that the use
of whole cells provided an easier and more efficient approach for enzyme retrieval. H. carterae, which is naturally
wall-less, is easily disrupted using a French pressure cell. A
typical PRK purification profile is presented in Table II.
Enzyme initially obtained from a 35 to 80% (NH4)2SO4
fractionation was desalted by exclusion chromatography
on a Sephadex G-50 column. The enzyme was further
purified by serial application of DE-52 chromatography
(Fig. 1A) followed by affinity chromatography with
agarose-Reactive Green 19 (Fig. 1B). This last step in PRK
isolation resulted in a highly enriched preparation (218
units mg21 protein) that was stored under N2 at 270°C in
the presence of glycerol, leupeptin, and DTT for at least 1
month without appreciable loss of activity.
All PRK enzymes observed to date are composed of two
to eight subunits. Subunit size for the H. carterae PRK
enzyme was electrophoretically resolved by 12% SDSPAGE. As seen in Figure 2, this chrysophytic enzyme is
composed of subunits that have a molecular mass of 53 6
1 kD. Additional minor protein bands were routinely observed in the enzyme preparation (Fig. 3). The sequence of
the amino terminus of the 53-kD protein verified a PRK
identity (Fig. 2; Table III). Because H. carterae Rubisco small
subunit protein showed significant sequence homology to
that of A. eutrophus (Boczar et al., 1989), and given the
clustered arrangement of these two genes in proteobacteria
(Gibson et al., 1990), we hypothesized that PRK and
Rubisco may have been moved into the chloroplast as a
laterally transferred cassette. However, antisera to A. eutrophus PRK failed to cross-react with this putative H.
carterae PRK subunit (data not shown).
To determine PRK holoenzyme size, affinity columnpurified protein was subject to Superose-6 flow performance liquid chromatography (Fig. 4). The molecular mass
of 214 6 12.6 kD observed in these experiments suggested
that H. carterae PRK is composed of four subunits.
Catalytic and Regulatory Properties
The kinetic characteristics of H. carterae PRK were analyzed using post-Reactive Green 19 affinity column-
Table II. H. carterae PRK purification procedure
PRK activity was monitored using a spectrophotometric assay. Enzyme units are micromoles of ADP per minute. Protein was quantified using
a protein assay kit (Bio-Rad) with BSA as a standard. Purification was determined using specific activity at each step compared with the specific
activity of the crude enzyme (0.7 unit mg21).
Step
Total Activity
Total Protein
Specific Activity
Purification
units
mg
units mg21
-fold
%
35– 80% (NH4)2SO4 fraction
86.8
86.80
1.0
100
Post-G-50 (Sephadex)
Post-DE-52 (anion exchange)
Post-Reactive Green 19 (affinity)
88.7
35.5
13.0
40.30
2.50
0.06
2.2
14.2
217.8
1.4
(from crude)
3.1
20.3
311.0
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Copyright © 1998 American Society of Plant Biologists. All rights reserved.
Units Recovered
100
41
15
Phosphoribulokinase in Marine Chromophytic Alga
325
Figure 2. H. carterae PRK protein eluted from the Reactive Green 19
affinity column was resolved on a 12% SDS-PAGE gel, transferred to
a PVDF membrane, and stained with Coomassie blue. The band (10
mg, lane 2) marked with an arrow was cut from the membrane,
sequenced at the amino terminus, and identified as PRK. Molecular
mass standards (lane 1) are a-lactalbumin (14.2 kD), trypsin inhibitor
(20.1 kD), carbonic anhydrase (29 kD), and ovalbumin (45 kD).
activity was not affected by NADH (Fig. 6A). In contrast,
DTT caused a significant increase in enzymatic activity
(Fig. 6B). Partially purified PRK (post-DE-52) was incubated with increasing concentrations of DTT before the
spectrophotometric assay was initiated with Ru5P. The
concentration of DTT in the spectrophotometric reaction
Figure 1. Chromatographic profile of H. carterae PRK fractions collected after Whatman DE-52 anion-exchange (A) and Reactive Green
19 affinity column chromatography (B). PRK activity was determined
using the spectrophotometric assay. Relative protein (post-DE-52)
and relative ATP concentration (post-Reactive Green 19) were determined by monitoring A280. The A280 for ATP concentration contained
less than 5% protein in each fraction.
purified enzyme. Classic Michaelis-Menten kinetics were
seen for both ATP and Ru5P, suggesting that no positive
cooperativity occurs between the enzyme and those substrates. Lineweaver-Burk plots resulted in an observed
Km(ATP) of 208 mm (Fig. 5A) and an observed Km(Ru5P) of
226 mm (Fig. 5B) for the enzyme.
To further assess whether H. carterae PRK was allosterically regulated by NADH (as found in many proteobacteria) or affected by a thioredoxin/Fd system (as seen in
cyanobacteria and terrestrial plants), the PRK was incubated at 4°C for at least 30 min in the presence of varying
concentrations of either NADH (0–1.0 mm) or DTT (0–30
mm). The enzyme was dialyzed against equilibration buffer
at 4°C for 2 h before incubation.
Partially purified PRK (post-Reactive Green 19) was incubated with increasing concentrations of NADH before
the radioactive assay was initiated with Ru5P. Enzyme
Figure 3. Coomassie blue- and silver-stained 12% SDSpolyacrylamide gels at various protein purification steps. Lysed H.
carterae cells containing PRK activity were fractionated with
(NH4)2SO4, passed through a Sephadex G-50 column, and further
purified on a DE-52 anion-exchange column followed by a ReactiveGreen 19 affinity column. The fractions with PRK activity are shown
after DE-52 anion-exchange (lane 2) and Reactive Green 19 affinity
column chromatography (lane 3). The PRK band is marked to the right
with an arrow. Molecular mass standards (lane 1) are trypsinogen (24
kD), carbonic anhydrase (29 kD), glyceraldehyde-3-phosphate dehydrogenase (36 kD), ovalbumin (45 kD), and BSA (66 kD).
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326
Hariharan et al.
Plant Physiol. Vol. 117, 1998
Table III. Alignment of amino N-terminal amino acid sequences of PRK from selected taxa
Residues in boldface letters represent the ATP-binding domain for oxygenic autotrophs. The shaded area indicates a conserved Cys. Dots
represent amino acid deletion.
Organism
Sequence
z
G
S
z
D
D
X
K E
K E
Ref.
Arabidopsis
Ice plant
Spinach
Wheat
Chlamydomonasreinhardtii
Selenastrum minutum A
Selenastrum minutum B
H. carterae
Odontella spp.
A
A
C
A
A
A
z
D
Q
V
K
G
G
G
G
Q
S
Q
E
D
L
L
E
E
E
Q
Q
Q
K
X
X
E
K
T
T
T
P
T
z
I
I V
I V
I V
I V
V V
V V
V V
z V V
P I V
z
z
z
z
z
z
z
L
z
I
I
I
I
I
I
G
G
G
G
G
G
G
I G
I G
L
L
L
L
L
L
L
V
V
A
A
A
A
A
A
A
A
A
A
A
A
A
A
A
A
A
A
D
D
D
D
D
D
D
D
D
S
S
S
S
S
Synechococcus sp. PCC 7942
Synechocystis sp. PCC 6803
Xanthobacter flavus
Rhodobacter sphaeroides
S K P
T T Q
M
V
z
L
S
S
D
D
I
K
R
R
K
K
L
L
I
I
I
I
I
I
V
V
V
V
A
A
T
V
G
G
G
G
D
D
S
S
S
S
S
S
V
V
H
Y
V
V
P
P
G
G
V
S
mixture was kept constant to ensure that the other enzymes
in the assay were fully active.
Amino-Terminal Amino Acid Sequencing
Proteobacterial PRK polypeptides have amino acid sequences that are completely divergent from those reported
for cyanobacteria and terrestrial plants (Porter et al., 1988;
Kossman et al., 1989). The amino termini of these two
kinase variants have signature sequences (Table III) that
identify each PRK type as either allosterically or thioredoxin/Fd regulated. Using Edman degradation, the aminoterminal 21 amino acids of H. carterae PRK were determined using protein obtained from the Reactive Green 19
affinity column. Its sequence demonstrates that the H. carterae polypeptide has a consensus ATP-binding domain that
shows almost perfect homology to that observed for PRK in
both cyanobacteria and terrestrial plants (thioredoxin/Fd
Figure 4. Determination of H. carterae PRK native size. Kav was
calculated as described in “Materials and Methods.”
G
G
G
G
G
C
C
C
C
C
G
G
G
G
G
K
K
K
K
K
S
S
S
S
S
T
T
T
T
T
F
F
F
F
F
S G C G K
S G C G K S T F
G
G
G
G
C
C
A
A
G
G
G
G
K
K
T
T
S
S
T
S
T
T
S
T
F
F
V
V
Horsnell and Raines (1991)
Michaelowski et al. (1992)
Roesler and Ogren (1988)
Raines et al. (1989)
Roesler and Ogren (1990)
Lin and Turpin (1992)
Lin and Turpin (1992)
This work
K. Weygrauch (1996) (GenBank
accession no. Y08610)
Wadano et al. (1995)
Su and Bogorad (1991)
Meijer et al. (199)
Gibson et al. (1990)
PRK). Also evident in this short sequence is a cysteinyl
residue (Cys-19) located within the nucleotide-binding domain. This amino acid serves as a regulatory disulfide in
thioredoxin/Fd PRK enzymes (Milanez et al., 1991). The
Edman method released a single amino acid at each cycle,
demonstrating homogeneity in the H. carterae PRK
polypeptide.
DISCUSSION
Enzyme Purification
Obtaining sufficient biomass provided an early challenge
in PRK isolation even though H. carterae was harvested at
h 6 a point in the 12-h light/12-h dark cell cycle when the
events of chloroplast biogenesis (transcription initiation,
photosynthetic capacity, protein accumulation) are maximal (Reith and Cattolico, 1985; Reynolds et al., 1993; Doran
and Cattolico, 1997). In the laboratory Heterosigma spp.
grow poorly in volumes of more than 1.5 L, and, like many
chromophytes, their cells fill with polysaccharide material
and other secondary metabolites as they enter later phases
of exponential cell growth (R.A. Cattolico, unpublished
data). This factor precluded cell harvest at high cell densities. Enzyme could be successfully retrieved from accumulated cells (40 L of culture was needed per enzyme purification procedure) when stored at 270° in the presence of
PMSF. This observation may be valuable to studies of PRK
in other unicellular chromophytes.
Treatment with 35 to 80% (NH4)2SO4 followed by Sephadex G-50 and ion-exchange chromatography provided a
small (20-fold) but effective initial step in H. carterae PRK
isolation. Most productive in the recovery of this enzyme
was the use of Reactive Green affinity chromatography
(300-fold purification). Triazine-based dyes that mimic coenzyme binding have been widely used as adsorbents for
the purification of dehydrogenases and kinases (Clonis and
Lowe, 1980). Although Cibacron Blue and Reactive Red
columns have been successfully used to isolate PRK from
both bacterial and eukaryotic sources (Siebert et al., 1981;
Ashton, 1984; Satoh et al., 1985; Porter et al., 1986; Serra et
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Copyright © 1998 American Society of Plant Biologists. All rights reserved.
Phosphoribulokinase in Marine Chromophytic Alga
327
would be rare events, and thus be taxonomically isolated.
Alternatively, such changes could occur frequently and, if
so, numerous variations in PRK holoenzyme structure
would be observed within close phylogenetic lineages.
Extensive analysis seems to verify a consistent octameric
PRK within all proteobacteria (see Tabita, 1980). Crystallographic analysis of R. sphaeroides shows the monomers to
be stacked in two planar tetramers (Roberts et al., 1995). In
contrast, sequence analysis of rRNA places the two genera
Anabaena (dimeric PRK) and Chlorogleopsis (hexameric
PRK) into sister groups within one of the eight phylogenetic clusters described for cyanobacteria (Wilmotte, 1994).
Both of these prokaryotes are filamentous and fix N2.
Given the phylogenetic proximity of these algae, one might
propose that the difference in PRK size results from a
simple rather than a complex evolutionary change.
Obviously, studies of other eukaryotic PRK enzymes are
needed. Although it is well established that chlorophytic
and chromophytic/rhodophytic taxa represent a major divergence in the evolution of autotrophic organisms, the
dimeric PRK structure routinely accepted for chlorophytes
(spinach, Scenedesmus, Selenastrum), and the tetrameric enzyme structure for chromophytes (Heterosigma), represent
an insufficient data set for good comparative analysis. Alternatively, one may argue that the shift in enzyme struc-
Figure 5. Rate of ADP production by partially purified H. carterae
PRK at various concentrations of ATP (0–1.5 mM, radioactive assay)
(A) and Ru5P (0–1 mM, spectrophotometric assay) (B). Insets show
Lineweaver-Burk plots of each data set. Km(ATP) 5 208 mM and
Km(Ru5P) 5 226 mM.
al., 1989), these columns did not bind the H. carterae enzyme efficiently.
Enzyme Size and Structure
As seen in Table II, PRK subunit size appears to vary
significantly among taxa. Terrestrial plant, chlorophytic
algae, and cyanobacterial PRK monomers (40 kD) are approximately 20% larger than those found in proteobacteria
(32 kD). The H. carterae monomer (53 kD) is more than 50%
greater in size than those in the proteobacterial cluster. It is
premature to speculate on the significance of this size
difference until the sequence of the H. carterae protein has
been completed and the PRK monomer size in a larger
number of non-chl b-containing plants has been assessed.
If the H. carterae enzyme is consistently found to be
tetrameric in future studies, then the question concerning
variability in quaternary structure among Calvin-cycle kinases must be addressed. It has been suggested by Meijer et
al. (1990) that changes in holoenzyme subunit number may
be related to differential conservation of domains critical to
subunit interaction. One could argue that these changes
Figure 6. Rate of ADP production by partially purified H. carterae
PRK at increasing concentrations of NADH (0–1 mM, radioactive
assay, SE less than 15%) (A) and DTT (0–30 mM, spectrophotometric
assay, SE less than 5%) (B).
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Copyright © 1998 American Society of Plant Biologists. All rights reserved.
328
Hariharan et al.
ture (i.e. dimeric u tetrameric state) may simply reflect a
means of regulating catalytic efficiency. Additional data
will also allow comparison of the evolutionary channeling
that has occurred for PRK (sequences and holoenzyme
structure) with that of Rubisco. At least for Rubisco, organelle coding site and ancestral proteobacterial identity
appear to be tightly correlated with either the chlorophyte
or chromophyte/rhodophyte lineages (for review, see
Delaney et al., 1995).
Microcompartmentation of Calvin-cycle enzymes occurs
in terrestrial plant and green algal chloroplasts. As many as
five enzymes, including PRK, can be found in large (500–
900 kD) arrays (Gontero et al., 1988; Süss et al., 1993; Wedel
et al., 1997) that theoretically enhance progression of catalytic intermediates from the active site of one enzyme to the
active site of another within the complex, enhancing enzymatic efficiency. No similar association of PRK in carboxysomes has been observed for either Thiobacillus neapolitanus
(Holthuijzen et al., 1986) or Chlorogleopsis fritschii (Marsden
et al., 1984). Suc-gradient analysis of disrupted H. carterae
cells suggests that the PRK of this alga may not exist in a
large complex. These data are consistent with observations
made by Mangeney et al. (1987), who failed to find PRK
within carboxysome-like structures in the cyanelles of Cyanophora paradoxa and Glaucocystis nostochinearum, which
had been immunocytochemically labeled with C. fritschii
PRK antiserum.
Enzyme Regulation
Protein sequencing of the H. carterae PRK amino terminus has revealed a distinct similarity between the kinase of
this chromophytic alga and those of cyanobacteria and
chlorophytes, but low sequence identity to proteobacterial
enzymes. All amino acids of the H. carterae ATP-binding
domain are homologous to those seen in cyanobacteria and
chlorophytic plants.
If H. carterae PRK is similar to enzymes found in oxygenic photoautotrophs, then one would expect that the
enzyme would be regulated by a thioredoxin/Fd cascade
rather than the allosteric control used by proteobacteria.
Sequence data (the presence of Cys-16) and the stimulatory
effect of DTT support this hypothesis. Like many thioredoxin/Fd PRK enzymes, H. carterae PRK quickly loses activity in an oxidized state, and compounds that have been
implicated in the alteration of proteobacterial PRK configuration (e.g. NADH [Fig. 6A]) appear to have no effect on
the DTT-activated catalytic efficiency of this enzyme.
Data show that H. carterae PRK displays hyperbolic kinetics similar to those seen for terrestrial plants and cyanobacteria for both ATP (Fig. 5A) and Ru5P (Fig. 5B). This
response is unlike the sigmoidal kinetics observed for proteobacteria, which indicate positive cooperativity for both
substrates (Abdelal and Schlegel, 1974). The Km values
calculated for H. carterae PRK were 208 mm (ATP) and 226
mm (Ru5P). A literature review of PRK function from photooxygenic species provides no clear answer to whether
Km(ATP) (53–1420 mm) or Km(Ru5P) (36–330 mm) values are
associated with holoenzyme structure or taxonomic affiliation (Gardemann et al., 1983; Satoh et al., 1985; Roesler
Plant Physiol. Vol. 117, 1998
and Ogren, 1990, Milanez et al., 1991; Su and Bogorad,
1991).
ACKNOWLEDGMENTS
We thank Carrine Blank for initiating these studies, Laurie
Connell for technical advice, William Hatheway for extensive support in this effort, and M. Kay Suiter for help in manuscript
preparation. R.A.C. dedicates this work to the memory of her
father, A.J. Cattolico.
Received September 30, 1997; accepted February 11, 1998.
Copyright Clearance Center: 0032–0889/98/117/0321/09.
LITERATURE CITED
Abdelal ATH, Schlegel HG (1974) Purification and regulatory
properties of phosphoribulokinase from Hydrogenomonas eutropha H 16. Biochem J 139: 481–489
Ashton AR (1984) An affinity label for the regulatory dithiol of
ribulose-5-phosphate kinase from maize (Zea mays). Biochem J
217: 79–84
Boczar BA, Delaney TP, Cattolico RA (1989) Gene for the
ribulose-1,5-bisphosphate carboxylase small subunit protein of
the marine chromophyte Olisthodiscus luteus is similar to that of
a chemoautotrophic bacterium. Proc Natl Acad Sci USA 86:
4996–4999
Clonis YD, Lowe CR (1980) Triazine dyes, a new class of affinity
labels for nucleotide-dependent enzymes. Biochem J 191: 247–251
Delaney T, Hardison L, Cattolico RA (1995) Evolution of plastid
genomes: inferences from discordant molecular phylogenies. In
C Sandgren, J Smol, J Kristiansen, eds, Chrysophytic Algae
Ecology, Physiology and Development. Cambridge University
Press, Cambridge, UK, pp 25–45
Doran E, Cattolico RA (1997) Photoregulation of chloroplast gene
transcription in the chromophytic alga Heterosigma carterae.
Plant Physiol 115: 773–781
Fuchs G (1986) CO2 fixation in acetogenic bacteria: variations on a
theme. FEMS Microbiol Rev 39: 181–213
Gardemann A, Stitt M, Heldt HW (1983) Regulation of spinach
ribulose-5-phosphate kinase by stromal metabolite levels. Biochim Biophys Acta 722: 51–60
Gibson JL, Chen J-H, Tower PA, Tabita FR (1990) The form II
fructose 1,6-bisphosphatase and phosphoribulokinase genes
form part of a large operon in Rhodobacter sphaeroides: primary
structure and insertional mutagenesis analysis. Biochemistry 29:
8085–8093
Gibson JL, Tabita R (1987) Organization of phosphoribulokinase
and ribulose bisphosphate carboxylase/oxygenase genes in Rhodopseudomonas (Rhodobacter) sphaeroides. J Bacteriol 169: 3685–
3690
Gontero B, Cárdenas ML, Ricard J (1988) A functional fiveenzyme complex of chloroplasts involved in the Calvin cycle.
Eur J Biochem 173: 437–443
Hallenbeck PL, Kaplan S (1987) Cloning of the gene for phosphoribulokinase activity from Rhodobacter sphaeroides and its expression in Escherichia coli. J Bacteriol 169: 3669–3678
Hart BA, Gibson J (1971) Ribulose 5-phosphate kinase from Chromatium sp. strain D. Arch Biochem Biophys 144: 308–321
Hartman H, Syyanen M, Buchanan BB (1990) Contrasting evolutionary histories of chloroplast thioredoxins f and m. Mol Biol
Evol 7: 247–254
Hemming A, Blotevogel KH (1985) A new pathway for CO2 fixation in methanogenic bacteria. Trends Biochem Sci 10: 198–200
Holmgren A (1985) Thioredoxin. Annu Rev Biochem 54: 237–271
Holthuijzen YA, van Breeman JFL, Kuenen JG, Konings WN
(1986) Protein composition of the carboxysomes of Thiobacillus
neapolitanus. Arch Microbiol 144: 398–404
Horsnell PR, Raines CA (1991) Nucleotide sequence of a cDNA
clone encoding chloroplast phosphoribulokinase from Arabidopsis thaliana. Plant Mol Biol 17: 183–184
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 1998 American Society of Plant Biologists. All rights reserved.
Phosphoribulokinase in Marine Chromophytic Alga
Kagawa T (1982) Isolation and purification of ribulose-5phosphate kinase from Nicotiana glutinosa. In M Edelman, RB
Hallick, N-H Chua, eds, Methods in Chloroplast Molecular Biology. Elsevier Press, Amsterdam, The Netherlands, pp 695–705
Kiesow LA, Lindsley BF, Bless JW (1977) Phosphoribulokinase
from Nitrobacter winogradskyi: activation by reduced nicotinamide adenine dinucleotide and inhibition by pyridoxal phosphate. J Bacteriol 130: 20–25
Kossmann J, Klintworth R, Bowien B (1989) Sequence analysis of
the chromosomal and plasmid genes encoding phosphoribulokinase from Alcaligenes eutrophus. Gene 85: 247–252
Laemmli UK (1970) Cleavage of structural proteins during the
assembly of the head of bacteriophage T4. Nature 227: 680–685
Lazaro JJ, Sutton CW, Nicholson S, Powls R (1986) Characterization of two forms of phosphoribulokinase isolated from the
green alga, Scenedesmus obliquus. Eur J Biochem 156: 423–429
Lin M, Turpin DH (1992) Purification and molecular and immunological characterization of a unique phosphoribulokinase
from the green alga Selenastrum minutum. Plant Physiol 98: 82–88
Loomis WF (1988) Four Billion Years: An Essay on the Evolution
of Genes and Organisms. Sinauer Associates, Sunderland, MA
Mangeney E, Hawthornthwaite AM, Codd GA, Gibbs SP (1987)
Immunocytochemical localization of phosphoribulose kinase in
the cyanelles of Cyanophora paradoxa and Glaucocystis nostochinearum. Plant Physiol 84: 1028–1032
Marsden WJN, Codd GA (1984) Purification and molecular and
catalytic properties of phosphoribulokinase from the cyanobacterium Chlorogleopsis fritschii. J Gen Microbiol 130: 999–1006
Marsden WJN, Lanaras T, Codd GA (1984) Subcellular segregation of phosphoribulokinase and ribulose-1,5-bisphosphate
carboxylase/oxygenase in the cyanobacterium Chlorogleopsis
fritschii. J Gen Microbiol 130: 2089–2093
McFadden BA, Tabita FR (1974) D-Ribulose-1,5-diphosphate carboxylase and the evolution of autotrophy. Biosystems 6: 93–112
McIntosh L, Cattolico RA (1978) Preservation of algal and higher
plant ribosomal RNA integrity during extraction and electrophoretic quantitation. Anal Biochem 91: 600–612
Meijer WG, Enequist HG, Terpstra P, Dijkhuizen L (1990) Nucleotide sequences of the genes encoding fructosebisphosphatase and phosphoribulokinase from Xanthobacter flavus H4–
14. J Gen Microbiol 136: 2225–2230
Michaelowski CB, DeRocher EJ, Bohnert H, Salvucci ME (1992)
Phosphoribulokinase from ice plant: transcription, transcripts
and protein expressions during environmental stress. Photosynth Res 31: 127–138
Milanez S, Mural RJ, Hartman FC (1991) Roles of cysteinyl residues of phosphoribulokinase as examined by site-directed mutagenesis. J Biol Chem 266: 10694–10699
Paulsen JM, Lane MD (1966) Spinach ribulose disphosphate carboxylase I. Purification and properties of the enzyme. Biochemistry 5: 2350–2357
Pharmacia Biotech (1993) Gel Filtration Principles and Methods,
Ed 6. Rahms i, Lund, Sweden, pp 6–13
Porter MA, Milanez S, Stringer CD, Hartman FC (1986) Purification and characterization of ribulose-5-phosphate kinase from
spinach. Arch Biochem Biophys 245: 14–23
Porter MA, Stringer CD, Hartman FC (1988) Characterization of
the regulatory thioredoxin site of phosphoribulokinase. J Biol
Chem 263: 123–129
Raines CA, Longstaff M, Lloyd JC, Dyer TA (1989) Complete
coding sequence of wheat phosphoribulokinase: developmental
329
and light-dependent expression of the mRNA. Mol Gen Genet
220: 43–48
Raven JA (1997) Putting the C in phycology. Eur J Phycol 32:
319–333
Reith M, Cattolico RA (1985) Chloroplast protein synthesis in the
chromophytic alga Olisthodiscus luteus. Plant Physiol 79: 231–236
Reynolds AE, McConaughy B, Cattolico RA (1993) Chloroplast
genes of the marine alga Heterosigma carterae are transcriptionally regulated during a light/dark cycle. Mol Mar Biol Biotech 2:
121–128
Rippel S, Bowien B (1984) Phosphoribulokinase from Rhodopseudomonas acidophila. Arch Microbiol 139: 207–212
Roberts DL, Runquist JA, Miziorko HM, Kim J-JP (1995) Crystallization and preliminary X-ray crystallographic analysis of
phosphoribulokinase from Rhodobacter sphaeroides. Protein Sci 4:
2442–2443
Roesler KR, Ogren WL (1988) Nucleotide sequence of spinach
cDNA encoding phosphoribulokinase. Nucleic Acids Res 14:
7192
Roesler KR, Ogren WL (1990) Chlamydomonas reinhardtii phosphoribulokinase sequence, purification, and kinetics. Plant Physiol
93: 188–193
Sainis JK, Harris GC (1986) The association of ribulose-1,5bisphosphate carboxylase with phosphoriboisomerase and phosphoribulokinase. Biochem Biophys Res Commun 139: 947–954
Satoh H, Okada M, Nakayama K, Murata T (1985) Purification of
ribulose 5-phosphate kinase and minor polypeptides of pyrenoid from the green alga Bryopsis maxima. Plant Cell Physiol 26:
931–940
Schäfer S, Götz M, Eisenreich W, Bacher A, Fuchs G (1989)
13
C-NMR study of autotrophic CO2 fixation in Thermoproteus
neutrophilus. Eur J Biochem 184: 151–156
Serra JL, Llama MJ, Rowell P, Stewart WDP (1989) Purification
and characterization of phosphoribulokinase from the N1-fixing
cyanobacterium Anabaena cylindrica. Plant Sci 59: 1–9
Siebert K, Schobert P, Bowien B (1981) Purification, some catalytic and molecular properties of phosphoribulokinase from Alcaligenes eutrophus. Biochim Biophys Acta 658: 35–44
Su X, Bogorad L (1991) A residue substitution in phosphoribulokinase of Synechocystis PCC 6803 renders the mutant lightsensitive. J Biol Chem 266: 23698–23705
Süss K-H, Arkona C, Manteuffel R, Adler K (1993) Calvin cycle
multienzyme complexes are bound to chloroplast thylakoid
membranes of higher plants in situ. Proc Natl Acad Sci USA 90:
5514–5518
Tabita FR (1980) Pyridine nucleotide control and subunit structure
of phosphoribulokinase from photosynthetic bacteria. J Bacteriol
143: 1275–1280
Taylor FGR (1992) The taxonomy of harmful marine phytoplankton. Giorn Bot Ital 126: 209–219
Wadano A, Kamata Y, Iwaki T, Nishikawa K, Hirahashi T (1995)
Purification and characterization of phosphoribulokinase from
the cyanobacterium Synechococcus PCC7942. Plant Cell Physiol
36: 1381–1385
Wedel N, Soll J, Paap BK (1997) CP12 provides a new mode of
light regulation of Calvin cycle activity in higher plants. Proc
Natl Acad Sci USA 94: 10479–10484
Wilmotte A (1994) Molecular evolution and taxonomy of the cyanobacteria. In DA Bryant, ed, The Molecular Biology of the Cyanobacteria. Kluwer Academic Publishers, Dordrecht, The Netherlands, pp 1–25
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 1998 American Society of Plant Biologists. All rights reserved.