A NOVEL ALKALINE PHOSPHATASE IN THE COCCOLITHOPHORE EMILIANIA HUXLEYI (PRYMNESIOPHYCEAE) AND ITS REGULATION BY PHOSPHORUS1

Abstract

Alkaline phosphatase (AP) plays an important role in the regeneration of bioavailable phosphate from organic compounds and allows phytoplankton growing in low inorganic phosphate environments to acquire phosphorus. We report the isolation, cloning, and initial characterization of the first AP (ehap1) in the coccolithophore Emiliania huxleyi (Lohm.) Hay and Mohler. This novel AP is a major form of AP released from the cell surface into the medium at late exponential and stationary phase of P-limited batch cultures but has no significant sequence similarity to other known APs. The cDNA sequence encodes a protein of roughly 94 kDa, while the processed active EHAP1 released from the cell surface is roughly 75 kDa. This difference is due to the cleavage of the signal sequence at its N-terminus and perhaps some truncation at its C-terminus. In response to variations in phosphate levels, the expression of ehap1 was found to correlate well with cellular AP activity. The ehap1 transcript was induced 4 h after phosphate depletion, increasing 1000-fold within a day, and was repressed rapidly upon phosphate addition. These results provide the basis for developing specific probes to study the expression of AP, and thus phosphate stress, of field populations.

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A NOVEL ALKALINE PHOSPHATASE IN THE COCCOLITHOPHORE
EMILIANIA HUXLEYI (PRYMNESIOPHYCEAE) AND ITS REGULATION BY
PHOSPHORUS1
Yan Xu2
Department of Ecology and Evolutionary Biology, Princeton University, Princeton, New Jersey 08544, USA
Thomas M. Wahlund
Department of Biological Sciences, California State University San Marcos, San Marcos, California 92096, USA
Liang Feng
Department of Molecular Biology, Princeton University, Princeton, New Jersey 08544, USA
Yeala Shaked3and Franc ¸ois M. M. Morel
Department of Geosciences, Princeton University, Princeton, New Jersey 08544, USA
Alkaline phosphatase (AP) plays an important
role in the regeneration of bioavailable phosphate
from organic compounds and allows phytoplankton
growing in low inorganic phosphate environments
to acquire phosphorus. We report the isolation,
cloning, and initial characterization of the first AP
(ehap1) in the coccolithophore Emiliania huxleyi
(Lohm.) Hay and Mohler. This novel AP is a major
form of AP released from the cell surface into the
medium at late exponential and stationary phase of
P-limited batch cultures but has no significant se-
quence similarity to other known APs. The cDNA
sequence encodes a protein of roughly 94kDa,
while the processed active EHAP1 released from
the cell surface is roughly 75kDa. This difference is
due to the cleavage of the signal sequence at its
N-terminus and perhaps some truncation at its
C-terminus. In response to variations in phosphate
levels, the expression of ehap1 was found to corre-
late well with cellular AP activity. The ehap1 tran-
script was induced 4h after phosphate depletion,
increasing 1000-fold within a day, and was re-
pressed rapidly upon phosphate addition. These
results provide the basis for developing specific
probes to study the expression of AP, and thus
phosphate stress, of field populations.
Key index words:
hophore; Emiliania huxleyi; Pacquisition; transcrip-
tional regulation
alkaline phosphatase; coccolit-
Abbreviations: 5PN, 50-nucleotidase; AP, alkaline
phosphatase; DOP, dissolved organic phosphorus;
Q-PCR, quantitative real-time PCR; RACE, rapid
amplification of cDNA ends; RT, reverse transcrip-
tion.
Phosphorus is often the main limiting nutrient for
primary production in freshwater and some marine
environments. Examples have been observed in Swed-
ish lakes, the eastern Mediterranean Sea, the northern
Red Sea, the central Atlantic, the Sargasso Sea, and the
oligotrophic Pacific gyre, where low phosphate con-
centrations often limit phytoplankton growth (Wu
et al. 2000, Stihl et al. 2001, Vidal et al. 2003, Krom
et al. 2004, Lomas et al. 2004, Bergstrom et al. 2005,
Thingstad et al. 2005). In these environments, dis-
solved organic phosphorus (DOP), which is usually
not directly available to phytoplankton, becomes the
most abundant P pool in surface waters (Karl and
Bjo ¨rkman 2002, Bjo ¨rkman and Karl 2003, Suzumura
and Ingall 2004). To acquire P from organic com-
pounds, the algae express extracellular phosphatases,
such as alkaline phosphatase (AP) and 50-nucleotidase
(5PN). These enzymes catalyze the hydrolysis of phos-
phate ester bonds in extracellular organic molecules,
permitting the cellular uptake of inorganic phosphate
(Pi) for metabolism.
AP enzymes are the best characterized extracellular
phosphatases. Their properties include low substrate
specificity and an alkaline pH optimum. The active site
of most APs contains a single Mg and two Zn atoms
(Coleman 1992). Many phytoplankton, such as dia-
toms, dinoflagellates, green algae, cyanobacteria, and
coccolithophoresexpress
(Kuenzler and Perras 1965, Dyhrman and Palenik
1997, Riegman et al. 2000, Stihl et al. 2001, Kruskopf
and Du Plessis 2004). AP can also be released into the
medium, as demonstrated in the diatom Phaeodactylum
APunder P-limitation
1Received 12 November 2005. Accepted 8 April 2006.
2Author for correspondence: e-mail yxu@princeton.edu.
3Present address: InterUniversity Institute for Marine Sciences,
Eilat 88103, Israel.
835
J. Phycol. 42, 835–844 (2006)
r 2006 by the Phycological Society of America
DOI: 10.1111/j.1529-8817.2006.00243.x

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tricornutum (Kuenzler et al. 1963, Chiaudani and Vighi
1982) and the green alga Chlamydomonas reinhardtii
(Quisel et al. 1996). AP enzymes from different phyto-
plankton species, and even isoenzymes within a given
organism, may have diverse substrate specificities,
physical and kinetic properties, and metal ion require-
ments (Rivkin and Swift 1980, Wagner et al. 1995, Karl
and Bjo ¨rkman 2002, Dyhrman and Palenik 2003).
Other than two reports for Synechococcus (Ray et al.
1991, Wagner et al. 1995), there are no reports de-
scribing the identification, cloning, and characteriza-
tion of an AP gene in photosynthetic phytoplankton.
AP expression in phytoplankton is regulated by ex-
tracellular and, in some cases, intracellular P concen-
trations, permitting an evaluation of the nutritional
status of natural phytoplankton populations based
upon AP activities (Dyhrman and Palenik 1999, Stihl
et al. 2001, Dyhrman et al. 2002, Vidal et al. 2003,
Lomas et al. 2004). In some studies, AP activities from
different oceanic regimes were measured in situ using
color-forming or fluorescent organic P compounds
(Hoppe 1983, Stihl et al. 2001, Sebastian and Niell
2004). Such bulk measurements provide only limited
information on the physiology of the ambient phyto-
plankton, however, because bacteria and zooplankton
may also contribute significantly to the total AP activity
(Hoppe 2003, Sebastian et al. 2004). Other methods,
such as single cell-based immunoassays or ELF (en-
zyme-labeled fluorescence) (Gonzalez-Gil et al. 1998,
Scanlan and Wilson 1999, Dyhrman and Palenik 2001,
Rengefors et al. 2001) allow investigators to identify
the organisms expressing AP, but provide no informa-
tion on enzyme hydrolytic rates. Some of these
analytical difficulties may be solved using an assay
that specifically targets AP in ecologically important
phytoplankton species. Development of such an assay
requires a sufficient characterization of AP in key or-
ganisms, and some data on its regulation as a function
of phosphate availability.
This study investigates an inducible AP from the
cosmopolitan marine coccolithophore Emiliania huxleyi
(Lohm.) Hay and Mohler. E. huxleyi has been shown to
express high AP activity and to possess a high-affinity P
uptake system, resulting in an exceptional phosphorus
acquisition capacity (Kuenzler and Perras 1965, Rieg-
man et al. 1992, 2000, Shaked et al. 2006). Analysis of
the kinetics of AP in E. huxleyi shows two distinct phases
(Riegman et al. 2000, Dyhrman and Palenik 2003),
suggesting the expression of more than a single type of
AP. In E. huxleyi strain CCMP 374, three phosphate-
regulated surface proteins are highly expressed under
P stress, and their expression level correlates with total
AP activity in the cells (Dyhrman and Palenik 2003).
However, none of these surface proteins have been
isolated, cloned, or characterized as yet.
Here, we report the first identification, cloning, and
initial characterization of an AP in E. huxleyi strain
CCMP 374. We also investigated the expression and
regulation of this enzyme in response to changes in
phosphorus concentration using quantitative real-time
PCR (Q-PCR). The results of this study demonstrate
that this AP responds rapidly to fluctuations in phos-
phorus levels in its environment, and thus, may play a
significant role in E. huxleyi in P-limited environments.
MATERIALS AND METHODS
Culture conditions. E. huxleyi strain CCMP374 was obtained
from the Provasoli-Guillard National Center for Culture of
Marin Phytoplankton in Maine, USA. Cells were grown in
0.2mm filtered and microwave-sterilized Gulf Stream seawa-
ter with the following filter sterilized components: 150mM
NaNO3, 1mM a-glycerophosphate, 59.3nM thiamine, and
Aquil trace metal mixture (Price et al. 1988/1989). Cultures
were incubated at 201 C under continuous light (80–
100mmol quanta?m?2?s?1), and cell growth was monitored
using a Multisizer II Coulter Counter (Beckman Coulter,
Hialeah, FL, USA).
AP activity assays. AP activity in the bulk medium was de-
termined by adding 50mL of 20mM p-nitro-phenylphos-
phate (p-NPP; in 1M Tris buffer at pH 8.2) into 1mL
sample to yield final concentrations of 1mM p-NPP and
20mM Tris at pH 8.2. The absorbance at 405nm was meas-
ured continuously for 2min using a Cary 100 UV-VIS Spec-
trophotometer (Varian, Vic., Australia). Enzyme activity was
computed from the linear regression of absorbance versus
time and then normalized to cell numbers. AP gel assays were
performed on a 4.8% non-denaturing gel using NBT/BCIP
staining. The gel was soaked in AP buffer (20mM Tris pH
8.0, 0.1mM MgCl2, 100mM NaCl) with 1% of 5-bromo-4-
chloro-3-indolyl phosphate (BCIP) and 0.01% of nitroblue
tetrazolium (NBT) in the dark at room temperature until
dark blue bands appeared. The gel was then washed with
water and fixed in 3% acetic acid (Manchenko 1994).
pH optimum measurement. The pH optimum of AP on cell
surface or in the filtrate (<0.22mm) was determined over
a range of pH 7.5–10 in seawater matrix. Buffers were
excluded as the application of various buffers over the ex-
periment’s wide pH range was found to influence the
enzyme’s pH optimum. For AP on cell surface, the intact
cells at late stationary phase were harvested by low-speed
centrifugation (15min at 8000g, 181 C) and were then resus-
pended in 4mL filtered seawater. For AP in the filtrate, the
cells were removed by a 0.2mm filter and the 2mL filtrate was
mixed with 2mL filtered seawater. The pH of the 4mL assay
solution was adjusted directly by adding 1M NaOH or HCl
before the activity assay and was measured again immediate-
ly afterward. When small pH shifts were noted, the reported
pH represents the average of both measurements. Enzyme
activity was measured as described above, except that the
final concentration of p-NPP was 0.5mM. It is worth men-
tioning that the pH optimum of the enzyme was influenced
by the p-NPP concentrations with lower pH optima at lower
substrate concentrations.
Protein purification. AP was partially purified from late sta-
tionary phase (15 days in stationary phase) culture supernat-
ants (2L) by low-speed centrifugation (15min at 8000g,
181 C) to pellet intact cells, followed by filtration through a
0.2mm filter to further remove cell debris. Activity assays of
the different fractions indicated that one half of the AP ac-
tivity remained in the culture filtrate. Proteins in this filtrate
were collected and concentrated using a 100kDa cut-off
ultra-filtration device (Millipore, Billerica, MA, USA). The
concentrated proteins were then purified by gel filtration on
a superose 6 column (Amersham Biosciences, Piscataway, NJ,
USA) in AP buffer. The fractions with the highest AP activity
were pooled and run on a non-denaturing gel, followed by an
in-gel activity assay. The same fractions were also run on a
12% SDS-PAGE gel, transferred onto a polyvinylidene fluo-
YAN XU ET AL.
836

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ride membrane, and the target protein band was excised for
N-terminal sequencing (Molecular Biology Department,
Princeton University). A BLASTsearch of the N-terminal se-
quence against the E. huxleyi EST sequence data base (Wah-
lund et al. 2004) yielded a single match. In order to obtain an
internal protein sequence directly from native gel, protein
purification was performed again as described above, with
the exception that the concentrated proteins were pre-incu-
bated with 5mM of dithiothreitol (DTT) for 1h at room tem-
perature before gel filtration. The DTTwas removed by gel
filtration against AP buffer. Owing to the low protein con-
centration, fractions with AP activity were pooled together
and concentrated with a 50kDa cut-off centricon (Millipore).
Proteins were loaded onto a native gel and an in-gel activity
assay performed. The band with AP activity was excised and
sent to ProtTech Inc. (Norristown, PA, USA) for peptide se-
quencing. In brief, the protein gel band was in-gel digested
with trypsin (modified sequencing grade; Promega, San Luis
Obispo, CA, USA), and the resulting peptide mixture was
sequenced by a Finnigan ion trap mass spectrometer LCQ
(ThermoFinnigan, San Jose, CA, USA) coupled on-line with
an HPLC system running a 75mM ID C18 column.
RNA isolation. Total RNA was extracted from cultures us-
ing TRIzol reagent (Invitrogen, Carlsbad, CA, USA) and an
RNeasy RNA extraction kit (Qiagen Inc., Valencia, CA, USA)
following the manufacturer’s instructions. The extracted
RNAs were treated with DNase I (1U, Invitrogen) to elim-
inate genomic DNA contamination.
Rapid amplification of cDNA 30end (30RACE) PCR amplifi-
cation. The cDNA was made using the first-strand cDNA syn-
thesis kit (Amersham Biosciences) with EhuxAda_T primer
(Table 1). Forward gene-specific primers (EhuxAP_1F and
EhuxAP_2F) were designed based on the EST sequence (Ta-
ble 1). The first round of amplification was performed with
the forward primer EhuxAP_1F and the reverse primer Eh-
uxAda_O in a total volume of 50mL containing 5mL of 10 ?
Pfx amplification buffer, 0.5mM of each deoxyribonucleoside
triphosphate, 0.3mM of each primer, 1mM of MgSO4, 5mL of
10 ? PCRxenhancer solution, 2.5U of PlatinumsPfx DNA
polymerase (Invitrogen), and 1mL of cDNA. The desired
PCR products were gel purified. The second round of nested
PCR was performed with the forward primer EhuxAP_2F
and the reverse primer EhuxAda_In in a total volume of
25mL containing 2.5mL of 10 ? Herculasesbuffer, 0.2mM
of each deoxyribonucleoside triphosphate, 0.6mM of each
primer, 4% of DMSO, 1U of HerculasesHotstart DNA
polymerase (Stratagene, La Jolla, CA, USA), and 1mL of
1000-fold diluted first-round PCR product. All thermal cy-
cling conditions and PCR reactions are given in Table 1. The
PCR products were purified and sequenced by Genewiz Inc.
(North Brunswick, NJ, USA), with several internal primers
designed by gene walking. Primers were from Integrated
DNA Technologies Inc. (Coralville, IA, USA).
Q-PCR amplification. Total RNAs were extracted and treat-
ed with DNase I as described above, and then quantified us-
ing the RiboGreen RNA Quantification Kit following exactly
the manufacturer’s instructions (Molecular probes, Invitro-
gen), and used to synthesize cDNA using the first-strand
cDNA synthesis kit (Amersham Biosciences) with a Not I-
d(T)18primer or a SuperScript III first-strand synthesis sys-
tem for RT-PCR (Invitrogen) with oligo(dT)20primer.
Q-PCR with gene-specific primers was used to determine
the induction and repression of ehap1 transcripts. The stand-
ard-curve quantification was used to quantify the exact level
of transcripts. The standard templates were prepared by first
amplifying ehap1 gene from E. huxleyi cDNA using gene-
specific primers EhuxAPRT1F and EhuxAPRT1R (Table 1),
second cloning into plasmids (TOPO TA Clonings, Invitro-
gen), third purifying from Escherichia coli cultures using QIA-
prep Spin Miniprep kit (Qiagen Inc.), and lastly quantifying
using the PicoGreensdsDNA quantification kit following the
manufacturer’s instructions (Molecular Probes, Invitrogen).
Serial dilutions of control template from 107to 10copies?mL?1
were used simultaneously in Q-PCR. The cycle number at
which the relative fluorescence of these serially diluted stand-
ards crossed the set threshold of amplification detection was
used for the Q-PCR analysis of mRNA extracted from exper-
imental cultures. Q-PCR analysis was performed on a
M ? 3000 (Stratagene) using the BrilliantsSYBRsGreen
QPCR Master Mix kit (Stratagene). The amplification of the
target gene was performed in triplicate for unknown samples,
and in duplicate for standards and no-RT controls. The reac-
tion mix contained 10mL of 2 ? master mix, 150nM of
EhuxAPRT1F and EhuxAPRT1R, 30nM of reference dye,
and cDNA synthesized from 20ng of total RNA. The thermal
cycling conditions of all runs are given in Table 1.
To evaluate induction of ehap1 transcription, the cells were
grown in medium containing 10mM inorganic PO4
other nutrients as described above. When the culture density
reached 3.2 ? 105cells?mL?1, cells were harvested by centri-
fugation at 8000g for 10min. The cells then were resuspended
in the same medium lacking phosphate (P-deplete) or alterna-
tively in medium containing Pi (10mM) but lacking nitrate
(P-replete). Cells were harvested every few hours for 1 day by
filtering onto 1mm polycarbonate membranes, freezing in liq-
uid nitrogen, and storing at ?801 C immediately. Cell densi-
ties and AP activities in the culture were measured at each
sample time point.
To evaluate repression of ehap1 transcription, cells were
transferred from P-replete medium to P-deplete medium as
described above and incubated for 12h. Cells were then di-
vided into two bottles, and 10mM Piwas added to one of them.
3?(Pi) and
TABLE1. Oligonucleotide primers, PCR reactions, and thermal cycling conditions used in this study.
PCR reactionF primer R primerTemplate Thermal cycling conditions
cDNA
synthesis
EhuxAda_T
(CCAGTGAGCAGAGTGACGA
GGACTCGAGCTCAAGC
TTTTTTTTTTTTTTTTT)
EhuxAda_O
(CCAGTGAGCAGAGTGACG)
EhuxAda_In
(GAGGACTCGAGCTCAAGC)
mRNA
30RACE
(Rnd1)
30RACE
(Rnd2)
EhuxAP_1F
(GGAGGGAGACTTCTGCTACG)
EhuxAP_2F
(CACCTTCATCAACACCGAGA)
EhuxAda_T-primed
cDNA
Purified 30
RACE PCR
30 ? (941 C, 30s; 571 C,
30s; 681 C, 3min)
571 C, 2min; 721 C,
40min; 29 ? (941 C, 15s;
571 C, 30s; 721 C, 3min)
40 ? (941 C, 30s; 561 C,
30s; 721 C, 30s)
Q-PCR EhuxAPRT1F
(AGCACATGTCGAACCCAA)
EhuxAPRT1R
(CGCCTCCACGAAGCAG)
Oligo(dT)-
primed cDNA
F, forward; R, reverse; Q-PCR, quantitative real-time PCR; RACE, rapid amplification of cDNA ends.
A NOVEL ALKALINE PHOSPHATASE IN EMILIANIA HUXLEYI
837

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Cells were harvested at 30min to 3h intervals for 12h, and
cell densities and AP activities were measured at each sample
time point.
RESULTS
Protein purification and identification. AP activity was
detected in the early stationary phase (day 5) in E.
huxleyi cultures supplemented with 1mM a-glycero-
phosphate (Fig. 1B). Based on the culture density at
this time point (330,000cells?mL?1) and cellular P
quotas (2.7–3.6fmol?cell?1, Shaked et al. 2006), we
estimated that ~99% of the P in the medium had
been taken up by the cells. Total AP activity in these
cultures increased about 7-fold from day 5 to day 15,
when theyreacheda
pNP?cell?1min?1(Fig. 1B, squares). AP activity
was also detected in the culture filtrate (<0.22mm)
and increased over time, suggesting release of the
enzyme from the cell surface (Fig. 1B, bars). More
than 50% of AP activity was detected in the culture
filtrate at day 21, which was then collected for further
purification.
Cultures were not pH-buffered. The pH changed
from 8.3 to 9.0 during the batch culture. The pH de-
pendence of AP activity, tested separately in the culture
filtrate and in whole cells (cell surface activity), showed
maximal values at pH59 ? 0.2 (Fig. 2). This value is
within the reported pH optima of 7.5–10.5 range for
AP (McComb et al. 1979, Maunders 1993). The cell
filtrate AP and the cell surface AP showed similar pH
maxima as well as enzyme kinetics (Shaked et al. 2006).
This indicates that AP released in the culture medium
exhibits characteristics similar to the AP associated with
cell surface, both showing an alkaline pH optimum.
maximum of~28pmol
Proteins (4100kDa) from the 21-day culture fil-
trate were fractionated by gel filtration and AP activity
eluted in two distinct peaks (Fig. 3). The first peak
(fractions 16–20) contained proteins larger than
800kDa, probably resulting from aggregation of small-
er proteins during purification. The second peak
(fractions 30–35) eluted proteins of about 160kDa.
SDS-PAGE (12%) of fraction 32 revealed two proteins
of approximately 115 and 75kDa (Fig. 4A). Although
the 75kDa protein band was much fainter than the
115kDa protein band, electrophoresis on a 4.8%
non-denaturing gel and in-gel assay indicated that
this smaller protein was AP (Fig. 4B). Therefore, the
N-terminus of the 75kDa protein was sequenced.
The N-terminal protein sequencing identified 10
amino acids of the 75kDa protein as: Ala Leu Ala Ser
Thr Asn Glu Tyr Leu Thr. A Blast search of the
106
105
104
103
A
B
40
30
20
10
00
5 10
Time (days)
15
AP activity
(pmol pNP · cell–1 · min–1)
cell number
(cells · mL–1)
20 25
FIG. 1. Growth curve and alkaline phosphatase (AP) activity
of Emiliania huxleyi cells grown with 1mM of a-glycerophosphate.
(A) Growth curve. (B) Induction of AP activity. Open squares:
total AP activity in the culture; bars: AP activity in the culture
filtrate (<0.22mM). The error bars represent the SD of the mean
of duplicate cultures.
Cells I
Cells II
Cells III
Filtrate I
Filtrate II
1.2
1.0
0.8
0.6
0.4
0.2
0.0
7.07.58.08.5 9.0
pH
Relative AP activity
9.510.0 10.5
FIG. 2. Relative activity of alkaline phosphatase (AP) on cell
surface and in culture filtrate at different pHs. For convenience,
the activities were normalized to the maximum activity at pH 9.0.
Three measurements were performed for AP on the cell surface
and two for AP in the culture filtrate.
Column fraction
10
0
50
100
150
200
250
15 2025 3035 40 4550
AP activity
(µmol pNP · min–1 · mL–1)
FIG. 3. Purification of proteins (4100kDa) from the culture
filtrate by gel filtration. Column fractions (0.5mL each) were
collected and assayed for alkaline phosphatase activity.
YAN XU ET AL.
838

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E. huxleyi EST data base identified a single EST pos-
sessing this N-terminal sequence. Sequence analysis of
this EST clone (clone ID 430277_G5_036) indicated
that the deduced N-terminal amino acids were within
the 50end of a gene we call ehap1. In order to obtain
the complete transcript of ehap1, we used 30-RACE (see
Methods), whose results yielded a 2.4kb fragment that
was cloned and sequenced (Fig. 5).
To verify that the cloned cDNA indeed encodes for
the AP that we identified in E. huxleyi, internal peptide
sequences were obtained from active protein excised
from non-denaturing gels. All 12 sequences obtained
fromthesetrypsin-digested
matched exactly the amino acid sequence deduced
from the cDNA nucleotide sequence (Fig. 4, under-
scored amino acids), confirming that the sequenced
gene encodes for a phosphatase, EHAP1.
Sequence. The ehap1 cDNA ORF was 2589bp long,
encodinga putativeprotein
93.8kDa (Fig. 5). The high GþC content (67%) was
similar to that described for other E. huxleyi coding
sequences (Wahlund et al. 2004). Approximately
6.1% and 7% of the amino acids are Asp and Glu, re-
spectively, contributing to the protein’s estimated pI
of 4.75. The protein also contains 6.8% of Pro with a
particularly high proportion (~17%) in the N-termi-
nal region. Translation of this cDNA sequence indi-
cated that 46 additional amino acids were located
upstream of the N-terminal protein sequence, and
in contrast to the rest of the protein, this region was
peptide fragments
of approximately
hydrophobic. Taken together, these data suggest that
the N-terminal region may be a transmembrane re-
gion that has to be cleaved off before release from the
cell surface.
Gel filtration indicated molecular masses of about
160kDa for the native EHAP1 and 75kDa for the de-
natured protein. The native form of this protein may
thus be a dimer, as is commonly observed in APs.
A search of GenBank against the deduced amino
acid sequence of EHAP1 did not identify a protein with
significant similarity to previously sequenced APs, nor
to any other known protein sequences. An interesting
exception was a hypothetical protein (gi33632062)
from Synechococcus sp. strain WH8102 that showed
23% identity (residues 221 through 643) to the
EHAP1 protein (residues 358 through 751). In addi-
tion, the EHAP1-predicted polypeptide sequence was
directly compared with APs from several eukaryotes
and prokaryotes, and no significant sequence homol-
ogy was found. Conserved sequences present in other
AP proteins were not present in EHAP1. The protein
folding predicted by 3D-PSSM (Kelley et al. 2000) re-
vealed no common structural motifs or domains be-
tween EHAP1 and other proteins in the data base.
Gene expression regulation by phosphate. In batch cul-
ture with 1mM a-glycerophosphate, AP activity was
not detectable until the late exponential phase when
P was depleted in the medium based on the calcula-
tion of P consumption by cells. Expression of the
ehap1 transcript was maintained at the minimal level
during the exponential phase (Fig. 6) and started to
increase, along with enzymatic activity, at the transi-
tion between the late exponential and the early sta-
tionary phase. The transcript level continued to
1
150
100
75
50
2
AB
34
FIG. 4. Proteins in fraction 32 (Fig. 2) were revealed by gel
electrophoresis. (A) SDS-PAGE (12%). Lane 1, molecular mass
standard (kDa); lane 2, proteins revealed by Coomassie blue
staining. (B) Non-denaturing gel (4.8%). Lane 3, result of in-gel
assay of alkaline phosphatase activity; lane 4, proteins revealed
by coomassie blue staining.
FIG. 5. Nucleotide sequence and deduced amino acid se-
quence of ehap1. N-terminal sequence is boxed and the matched
internal peptide sequences are underscored.
10
9
8
7
6
0
4.0 4.55.0
Time (days)
5.5 6.06.5
5
10
15
0
2
1
3
4
105 cell number
(cells · mL–1)
AP activity
(pmol pNP · cell–1 · min–1)
103 mRNA copy number
(/40 ng total RNA)
A
B
FIG. 6. Growth curve, alkaline phosphatase (AP) activity, and
ehap1 transcript level of Emiliania huxelyi cells over time during
the transition between exponential and stationary phases. Cells
were grown with 1mM of a- glycerophosphate. (A) Growth curve.
(B) Open squares, total AP activity in the culture (right Y-axis);
closed squares, ehap1 transcript abundance calculated as copy
numbers in 40ng of total RNA (left Y-axis). The error bars rep-
resent the SD of the mean of triplicate measurements of mRNA
copy numbers.
A NOVEL ALKALINE PHOSPHATASE IN EMILIANIA HUXLEYI
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increase for more than a day after cells entered the
stationary phase (Fig. 6). In a short-term induction
experiment, the ehap1 transcript level did not in-
crease until 4h after transfer into P-deplete medium
and transcript abundance increased about 1000-fold
during the following 4–24h (Fig. 7). Meanwhile, the
ehap1 transcript level was maintained at the original
level after transfer in P-replete medium. AP activity in
cells was only detectable in P-deplete medium after
about 12h. The lag between the increase of transcript
level and the detection of enzymatic activity probably
reflects in part de novo protein synthesis and local-
ization, as well as the relatively low sensitivity of the
enzyme assay.
The repression of ehap1 transcript was also tested.
After cells were transferred from P-replete medium to
P-deplete medium for 12h, the ehap1 transcript in-
creased ~500-fold from the minimum level (0h in
Fig. 8A). Upon phosphate addition, a significant de-
crease in the ehap1 transcript was detected within 1h,
followed by a gradual decrease to the minimum level at
6h. Without phosphate addition, ehap1 transcript level
remained relatively constant over 2h, and then in-
creased about 6-fold by 6h. In contrast, AP activity did
not decrease significantly for 6h after phosphate ad-
dition, whereas it increased continuously in P-deplete
cells (Fig. 8B). The contrast between the RNA and the
AP activity responses upon P addition probably reflects
the fact that it takes a few days for cells to degrade ex-
tracellular AP. Nonetheless, protein synthesis was sup-
pressed by P addition as there was no further increase
in enzymatic activity. Overall, there was a good corre-
lation between the transcript level of ehap1 and the
whole cell enzyme activity.
DISCUSSION
The identification of the protein whose gene we
have cloned as a phosphatase has been firmly estab-
lished by the alignment of internal peptide sequences
from a protein with phosphatase activity with the de-
duced amino acid sequence from ehap1 cDNA. Fur-
thermore, the regulation of ehap1 expression by
phosphate is consistent with the regulation pattern of
phosphatases in other organisms (Rivkin and Swift
1979, Dyhrman and Palenik 2003), and the RNA ex-
pression level is well correlated with total phosphatase
activity in the culture. The enzyme is most likely an AP
(rather than a 50nucleotidase), given that known AP
substrates, PNPP, BCIP, and DiFMUP (6,8-difluoro-4-
methylumbelliferyl phosphate), are active substrates
for EHAP1, the enzyme has an alkaline pH optimum,
and ehap1 expression is repressed rapidly by Pi(Am-
merman and Azam 1985).
This enzyme, EHAP1, is a novel protein without any
similarity to other known APs. This result was unex-
pected as the amino acid sequences of known APs ap-
pear to have been well conserved during evolution,
especially in the region of their active sites (Murphy
80
60
40
20
0
1.5
1.0
0.5
0.0
0510
Time (hours)
1520 25
103 mRNA copy number
(/40 ng total RNA)
AP activity
(pmol pNP · cell–1 · min–1)
FIG. 7. Induction of ehap1 transcript by P limitation. ehap1
transcript abundance was calculated as copy number in 40ng of
total RNA. Closed circles, transcript level in P-deplete culture
(left Y-axis); open circles, transcript level in P-replete culture (left
Y-axis); and squares, total alkaline phosphatase activity in P-de-
plete culture (right Y-axis). The error bars represent the SD of
the mean of triplicate measurements of mRNA copy numbers.
50
40
30
10
5
0
012345
Time (hours)
Time (hours)
670
0.0
0.1
0.2
0.3
0.4
0.5
1234567
103 mRNA copy number
(/40 ng total RNA)
AP activity
(pmol pNP · cell–1 · min–1)
AB
FIG. 8. (A) Repression of ehap1 transcript by inorganic phosphate. The ehap1 transcript abundance was calculated as copy number in
40ng of total RNA. Closed circles, AP activity in P-deplete culture; open circles, transcript level in P-replete culture. (B) Total alkaline
phosphatase activity. Closed circles, transcript level in P-deplete culture; open circles, AP activity in P-replete culture. The error bars
represent the SD of the mean of triplicate measurements of mRNA copy numbers.
YAN XU ET AL.
840

Page 7

et al. 1995). In marine phytoplankton, putative AP
genes in Thalassiosira pseudonana and phoA genes in sev-
eral Trichodesmium species are also similar to AP genes
characterized in GenBank (Orchard et al. 2003). In
Synechococcus sp. strain PCC 7942, which possesses two
AP genes, phoV is similar to other APs while phoA is not
(Ray et al. 1991, Wagner et al. 1995). Our finding of a
novel AP in E. huxleyi is consistent with the notion that
the genomes of marine phytoplankton may contain
many novel genes (Armbrust et al. 2004, Venter et al.
2004, Lane et al. 2005, Montsant et al. 2005). In E.
huxleyi, more than 50% of the EST sequences from its
data base show no significant similarity to known pro-
teins in GenBank (Wahlund et al. 2004). Consistent
with our result, no genes similar to known APs have
been found in studies of differences in expression pro-
files between P-replete and P-deplete E. huxleyi cultures
(Wahlund et al. 2004, Nguyen et al. 2005).
A gene sequence (prp1) highly similar to ehap1 has
been found in another strain of E. huxleyi (strain
CCMP1516; Landry et al. 2006). The alignment of
the amino acid sequences revealed changes in only five
residues between these two strains. The cDNA se-
quences were identical, except for the corresponding
changes. We have tested the induction of putative
ehap1 transcript in nine other strains of E. huxleyi iso-
lated from different regions of the ocean using gene-
specific primers EhuxAPRT1F and EhuxAPRT1R.
The transcript level increased by 102- to 105-fold in
all strains tested 24h after cells were transferred from
P-replete to P-deplete medium, which indicates that
ehap1 is highly conserved in E. huxleyi (Y. Xu, unpub-
lished data). This is consistent with previous studies of
E. huxleyi that have revealed very little inter-clonal var-
iation in the DNA sequences coding for either the small
subunit rRNA gene, the untranscribed RUBISCO
rbcL-rbcS spacer region, or the transcribed GPA region
(Medlin et al. 1994, Schroeder et al. 2005).
The EHAP1 precursor (93.8kDa) is markedly larg-
er than its mature form (75kDa). This could be partly
explained by the potential cleavage site in the precur-
sor (between Pro 16 and Glu 17), which would release
a putative signal peptide (Bendtsen et al. 2004). Com-
parison of the N-terminal sequence data against the
deduced amino acid sequence suggests that an addi-
tional cleavage may occur between Leu 46 and Ala 47.
After the predicted cleavage, the size of the processed
protein would be 88.9kDa, which is similar to the size
of PRP1 (89.3kDa) in the EDTA-solubilized fraction
from strain CCMP1516 (Landry et al. 2006). However,
the processed protein would still be larger than that
measured by gel electrophoresis for the isolated pro-
tein. Some possible explanations include alternative
splicing of mRNA during post-transcriptional process-
ing and proteolytic cleavage during post-translational
modification. There are two possible alternatives for
proteolytic cleavage of the C-terminus: (1) regulated
proteolytic cleavage by specific proteases for the
release of membrane-bound proteins (Sisodia 1992,
Destrooper et al. 1993, Massague and Pandiella 1993,
Parkin et al. 2004); and (2) unregulated proteolytic
cleavage by proteases induced under nutrient depri-
vation (Berges and Falkowski 1996, 1998, Bidle and
Falkowski 2004). Further experiments are required to
resolve this type of question.
There is little information available bearing on the
question of how EHAP1 is anchored to the cell mem-
brane and then released into the medium. Some ex-
ternal APs from aquatic plants and animal tissues
(Wong and Low 1992, 1994, Morita et al. 1996) are at-
tached to the cell membrane by a GPI anchor (glyco-
sylphosphatidylinositol anchor). The release of these
proteins often involves the action of a GPI-specific
phospholipase (Takami et al. 1992, Morita et al. 1996,
Yoda et al. 2000, Ikezawa 2002). However, the lack of
hydrophobicity in the C-terminus of EHAP1 suggests
that it is not a GPI-anchored protein (Kronegg and
Buloz 1999). Similarly, a GPI anchor was not identified
in AP from Prorocentrum minimum (Dyhrman 2005).
AP is normally induced shortly after the depletion of
phosphate from the medium and before the growth
rate declines. It thus appears that the phosphate con-
centration in the medium may be an important factor
in regulating the expression of AP (Cembella et al.
1984). The external P concentration required to in-
duce AP activity in E. huxleyi is extremely low (nM
range; Shaked et al. 2006). Our results showing that
the induction of ehap1 expression was delayed for sev-
eral hours after transfer into P-deplete medium pre-
sumably reflects the slow consumption of the stored
intracellular P. According to Shaked et al. (2006),
the cellular P quota in P-replete medium (4.8–
5.7fmol?cell?1) is higher than that in low-P medium
(2.7–3.6fmol?cell?1). Based on the cell density (not
shown) at each time point in Fig. 7, we estimated that
8–14h were needed to consume the stored intracellu-
lar P, which is consistent with the observation that the
rapid increase of ehap1 transcript initiated around 10h.
The internal P pool may thus also be an important
regulatory factor for AP in E. huxleyi.
At the RNA level, the time required for E. huxleyi to
induce ehap1 expression upon the depletion of P in the
medium is comparable with that reported for phoA in
Synechococcus sp. strain PCC 7942 (Ray et al. 1991).
However, following P-addition to P-starved cells, the
reduction in ehap1 transcript levels is much slower than
that of phoA. The induction time is also comparable to
that of genes described for other nutrient acquisition
systems, such as the nitrate reductase gene in Dunali-
ella tertiolecta (Song and Ward 2004), narB (nitrate red-
uctase) and nirA (nitrite reductase) in Synechococcus sp.
Strain WH 8103 (Bird and Wyman 2003), and twca1
(carbonic anhydrase) in Thalassiosira weissflogii (Lane
and Morel 2000). However, the level of induction of
ehap1 is significantly higher than reported for any of
the above genes, and the induction period is longer.
This is consistent with the extremely high AP activity
observed in E. huxleyi. At the protein level, the time
required for the induction of AP activity in E. huxleyi
(12h) is similar to that in C. reinhardtii (Quisel et al.
A NOVEL ALKALINE PHOSPHATASE IN EMILIANIA HUXLEYI
841

Page 8

1996), but apparently longer than the induction of CA
activity in T. weissflogii (5h) (Lane and Morel 2000).
This apparent difference may simply reflect the great-
er sensitivity of the CA assay, however.
In summary, we have identified and partially char-
acterized a novel AP, with no similarity to previously
known APs in the marine coccolithiphore E. huxleyi.
The dissimilarity between EHAP1 and other APs and
the rapid regulation of ehap1 upon changes in phos-
phorus concentration suggest that this enzyme may be
a valuable target for field studies. In many coastal and
oceanic regions, E. huxleyi often forms extensive blooms
(Brown and Yoder 1994, Iglesias-Rodriguez et al.
2002, Tyrrell and Merico 2004) and the role that mac-
ronutrients play in triggering these blooms is a matter
of debate (Tyrrell and Merico 2004, Lessard et al.
2005). The NO3
cator for the nutritional status of E. huxleyi in the field,
given that some other forms of nitrogen and phospho-
rus are available (Palenik and Henson 1997, Shaked
et al. 2006) and the physiology of the field population
may be changing during bloom development. The
measurement of the expression level of ehap1, in con-
cert with AP activity assay, may provide a specific indi-
cator of phosphate stress in E. huxleyi in the field.
?:PO4
3?ratio may not be a valid indi-
We thank T. G. Doak for technical assistance with primer de-
sign and B. P. Palenik for helpful discussions. We thank Y. Shi
for the use of his FPLC and B. Ward for the use of her Q-PCR
system. This work was supported by the Center for Environ-
mental BioInorganic Chemistry (NSF grant 0221978) and
NSF grant 0351499 to F. M. M. Morel.
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