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First published online July 10, 2003; 10.1104/pp.103.020008 Plant Physiology 132:1840-1848 (2003) © 2003 American Society of Plant Biologists Subcellular Targeting of Nine Calcium-Dependent Protein Kinase Isoforms from Arabidopsis1Department of Cell Biology, The Scripps Research Institute, La Jolla, California 92037 (C.D., B.H., S.M.R., J.F.H.); Biology Department, Washington University, St. Louis, Missouri 63130–4899 (A.I., B.G.P.); Department of Plant Biology, University of New Hampshire, 46 College Road, Durham, New Hampshire 03824 (E.M.H.); and Botany Department, University of Florida, Gainesville, Florida 32611–8526 (A.C.H.)
Calcium-dependent protein kinases (CDPKs) are specific to plants and some protists. Their activation by calcium makes them important switches for the transduction of intracellular calcium signals. Here, we identify the subcellular targeting potentials for nine CDPK isoforms from Arabidopsis, as determined by expression of green fluorescent protein (GFP) fusions in transgenic plants. Subcellular locations were determined by fluorescence microscopy in cells near the root tip. Isoforms AtCPK3-GFP and AtCPK4-GFP showed a nuclear and cytosolic distribution similar to that of free GFP. Membrane fractionation experiments confirmed that these isoforms were primarily soluble. A membrane association was observed for AtCPKs 1, 7, 8, 9, 16, 21, and 28, based on imaging and membrane fractionation experiments. This correlates with the presence of potential N-terminal acylation sites, consistent with acylation as an important factor in membrane association. All but one of the membrane-associated isoforms targeted exclusively to the plasma membrane. The exception was AtCPK1-GFP, which targeted to peroxisomes, as determined by covisualization with a peroxisome marker. Peroxisome targeting of AtCPK1-GFP was disrupted by a deletion of two potential N-terminal acylation sites. The observation of a peroxisome-located CDPK suggests a mechanism for calcium regulation of peroxisomal functions involved in oxidative stress and lipid metabolism.
Calcium-dependent protein kinases (CDPKs) are Ser/Thr protein kinases that are only found in plants and some protozoans. They are composed of a variable N-terminal domain, a catalytic domain, an auto-inhibitory region, and a calmodulin-like domain (Harper et al., 1991  ; Suen and Choi,
1991; Huang et. al.,
1996; Cheng et al.,
2002; Hrabak et al.,
2003). CDPKs are activated upon binding calcium to their
calmodulin-like domain, which makes them effective switches for the
transduction of calcium signals in plant cells.
Calcium signaling has emerged as a central mechanism to regulate responses
to environmental stimuli such as cold, salt stress, and drought (for review,
see Knight and Knight, 2001
There are 34 CDPK genes in Arabidopsis. Although these kinases all have
strongly conserved catalytic and calmodulin-like regulatory domains (Harmon et
al., 2000
There is evidence for CDPKs in multiple subcellular locations, including
the plasma membrane (e.g., Schaller et
al., 1992 Here, we provide evidence for the potential subcellular locations of nine additional Arabidopsis isoforms, based on localization of green fluorescent protein (GFP) fusions. Our results delineate three additional targeting patterns for the Arabidopsis CDPK family, with membrane association observed for all seven isoforms harboring potential N-terminal myristoylation sites. Interestingly, plasma membrane association was observed for six of these seven isoforms. The exception was AtCPK1-GFP, which was found associated with peroxisomes. These results demonstrate that in Arabidopsis, different CDPKs are targeted to specific subcellular locations, providing the potential for isoform-specific differences in regulating various cellular functions. In the case of AtCPK1, our results provide the first evidence for the potential role of a CDPK in regulating peroxisomal functions such as lipid metabolism and oxidative stress.
Subcellular Locations of AtCPK-GFP Fusion Proteins in Arabidopsis Root Tips Potential subcellular locations for nine CDPKs were determined by expression of AtCPK-GFP fusions in stable transgenic plants. Isoforms AtCPKs 1, 3, 4, 7, 8, 16, 21, and 28 were chosen as a starting point for a complete survey of all 34 Arabidopsis isoforms, because these genes were available as full-length cDNAs at the initiation of the project. Each kinase was engineered with a C-terminal GFP and was expressed in plants under the control of a cauliflower mosaic virus 35S promoter. The nine isoforms revealed three different subcellular distribution patterns, as detected by fluorescence confocal microscopy of cells located near the root tip (Fig. 1).
The subcellular distribution of GFP alone is shown as a control in Figure 1A. These images provide a reference for cytosolic and nuclear localization patterns, as previously shown (e.g., Haseloff et al., 1997 ). For each CPK-GFP construct, multiple transgenic lines were
examined, and images were analyzed for cells showing both high and low levels
of expression. The patterns shown were observed consistently, even at the
lower limits of detection in weakly fluorescent cells. This supports the
contention that the observed localization patterns are not an artifact of
extreme overexpression but rather reflect realistic targeting potentials for
endogenous enzymes.
Imaging of isoforms AtCPK3-GFP and AtCPK4-GFP revealed distribution
patterns similar to that of free GFP, including nuclear fluorescence in most
cells (Fig. 1, C and D). This
suggests that both isoforms are primarily soluble enzymes with the potential
to target to the nucleus, although neither AtCPK3 nor AtCPK4 contains nuclear
localization signals when analyzed by ProSite
(Falquet et al., 2002
The six GFP fusions for isoforms AtCPKs 7, 8, 9, 16, 21, and 28 were all observed in a thin layer at the periphery of the cells, consistent with a plasma membrane location. Moreover, all of these GFP fusions cofractionated with the microsomal membrane fraction (Fig. 2; AtCPK7-GFP is shown as a representative example). AtCPK1-GFP was primarily seen in small, often spherical organelles approximately 0.5 to 1.5 µm in diameter, which is the typical size of mitochondria and peroxisomes (Fig. 1B). Consistent with an association with an organellar membrane, AtCPK1-GFP cofractionated with the microsomal membrane fraction, as shown by western-blot analysis in Figure 2.
To determine the identity of AtCPK1-GFP-associated organelles, we conducted covisualization experiments with markers for mitochondria and peroxisomes. For covisualization with mitochondria, we used the mitochondrion-specific fluorescent dye MitoTracker Red. The projected three-dimensional image in Figure 3A shows very little overlap between the MitoTracker and GFP signals. This result supports a non-mitochondrial location for AtCPK1-GFP.
To test whether AtCPK1-GFP was colocalized with peroxisomes, we first
compared plant cells expressing either AtCPK1-GFP or a peroxisome targeted
GFP. A modified GFP was targeted to peroxisomes by the addition of a
C-terminal sequence KSRM (GFPrs-KSRM;
Trelease et al., 1995 To further verify a peroxisome location, we conducted covisualization experiments in which peroxisomes and AtCPK1 were labeled with two spectrally distinct GFPs. For the experiment shown in Figure 3, D through F, AtCPK1 was labeled with a "sharp-green" GFP (GFPsg) with an excitation spectrum peaking at 398 nm. The peroxisome-targeted GFPrs-KSRM was constructed with a "red-shifted" GFP (GFPrs) with an excitation peak at 488 nm (similar to EGFP, which was used for all other CDPK-GFP fusions). Both GFPs retained the same emission peak at 511 nm. Controls showed that each type of GFP could be uniquely visualized using the excitation filters and exposure times used in this study. When AtCPK1-GFPsg and GFPrs-KSRM were co-expressed in stably transformed plants, their fluorescence images overlapped with each other (Fig. 3, D–F). Equivalent results were obtained with a second set of constructs in which the GFPsg and GFPrs tags were switched with respect to the labeling of CPK1 and a peroxisome-targeted GFP (not shown). This colocalization analysis strongly supports a peroxisomal location for AtCPK1-GFP. A final line of evidence for a peroxisomal location of AtCPK1-GFP is the distinct "torus" morphology that could be observed in high-magnification images, such as Figure 3D. This torus or "doughnut" morphology has previously been noted for images of plant peroxisomes (e.g. Cutler at al, 2000; http://deepgreen.stanford.edu).
As predicted for many CDPKs, AtCPK1 has two potential acylation sites at
its N-terminal end. To investigate whether these acylation sites have a role
in peroxisome targeting, they were removed by swapping the seven N-terminal
amino acid residues (MGNTCVG-P8) of AtCPK1 with the four N-terminal
residues from AtCPK12 (MANK-P5). The rationale was to swap
N-terminal extensions defined by the sequences located upstream from the first
Pro in each CDPK (positions 8 and 5, respectively). This sequence exchange
removed all potential acylation sites, namely the Gly in position 2 (for
myristoylation) and the Cys in position 5 (for palmitoylation). The resulting
mutant construct, referred to as AtCPK1-
To confirm the disruption of peroxisome localization, we fractionated
membranes and soluble proteins and performed a western-blot analysis. The
wild-type AtCPK1-GFP was more abundant in the membrane fraction, whereas the
mutant AtCPK1-
CDPKs Are Located in Many Subcellular Locations
To understand the isoform-specific functions of all 34 Arabidopsis CDPKs,
it will be necessary to delineate their subcellular locations, tissue
specificity, substrate specificities, and biochemistry. Here, we provide a
comparison of subcellular targeting properties for nine AtCPK isoforms. Our
results provide evidence for: (a) a cytosolic and nuclear location for
isoforms AtCPKs 3 and 4 (Fig. 1, C and
D), (b) a plasma membrane location for isoforms AtCPKs 7, 8, 9,
16, 21, and 28 (Fig. 1,
E–J), and (c) a peroxisome membrane location for isoform
AtCPK1 (Figs. 1B and
4). Together with evidence for
an ER location of AtCPK2 (Lu and Hrabak,
2002
This survey includes at least two CDPKs from each of the four subfamilies
(Hrabak et. al., 2003
Although our survey clearly identifies isoform-specific differences in
targeting potentials, we did not verify the actual locations of endogenous
(untagged) kinases. Our approach was limited to a survey of GFP-tagged
isoforms, expressed using a 35S promoter, and imaged in root tip cells.
Potential differences between locations of GFP-tagged and endogenous CDPKs
could result from several factors, including the following: (a) The addition
of a C-terminal GFP-tag may interfere with proper targeting, for example, by
disrupting a targeting signal or a protein-protein interaction. (b) The
imaging of isoforms ectopically expressed in root tip cells may fail to reveal
targeting potentials unique to the specific cell type, such as a guard cell or
pollen tube. (c) The environmental conditions under which the roots were grown
and imaged may have failed to reveal a conditional targeting potential, such
as seen for Mc-CPK1, which appears to translocate from the plasma membrane to
the nuclei of ice plant (Mesembryanthemum crystallinum) epidermal
cells only after exposure to salt stress
(Patharkar and Cushman, 2000
Of the seven membrane-associated CDPKs examined here, AtCPK1-GFP was the only one that was not associated with the plasma membrane. Two lines of evidence support the contention that AtCPK1-GFP is associated with peroxisomes. First, imaging of GFP-tagged AtCPK1 indicated that AtCPK1-GFP is associated with 0.5- to 1.5-µm organelles, a size most consistent with peroxisomes and mitochondria. These organelles were shown not to be mitochondria by covisualization experiments with a mitochondrion-specific dye (Fig. 3A). In contrast, colocalization experiments with a peroxisomal marker showed a high degree of overlap (Fig. 3, B–F). Second, when AtCPK1-GFP fluorescence was examined at high magnification, the spherical bodies were frequently observed to have a doughnut morphology. This same morphology was observed with a marker (GFPrs-KSRM) targeted to the lumen of peroxisomes. A similar morphology has also been noted for other peroxisome-targeted proteins (http://deepgreen.stanford.edu), including a common peroxisomal marker (catalase) tagged with GFP.
Although we have not excluded the possibility that AtCPK1-GFP is located
within the peroxisome, occurrence at the cytosolic surface is considered more
likely. This is based on our observation that peroxisome localization was
disrupted by removal of the N-terminal acylation sites of AtCPK1
(Fig. 4). To our knowledge,
there is no precedent for the involvement of an N-terminal myristoyl or
palmityl group in peroxisomal import. Thus, the simplest explanation is that
acylation potentiates an AtCPK1 peroxisome membrane interaction analogous to
the membrane interactions of CDPKs localized to the ER or plasma membrane
(Martin and Busconi, 2000
Most CDPKs have predicted N-terminal acylation sites that could promote
membrane association. This prediction has been experimentally confirmed for
three isoforms: OsCPK2, LeCPK1, and AtCPK2
(Martin and Busconi, 2000
At present, the majority of membrane-associated CDPKs have been found
localized to the plasma membrane (e.g. AtCPKs 7, 8, 9, 16, 21, 28, LeCPK1, and
OsCPK2). However, two of the Arabidopsis isoforms are now known to associate
with endomembranes, with AtCPK1 associated with peroxisomes (this study) and
AtCPK2 with ER (Lu and Hrabak,
2002 In the context of isoform-specific targeting of CDPKs, it is noteworthy that the N-terminal acylation sites are located in highly variable N-terminal domains. Because acylation is only one of several features contributing to membrane association, we speculate that other sequences within the N-terminal domains may provide the basis for isoform-specific protein-protein interactions. Even among the isoforms known to target to the plasma membrane, there is no obvious conserved sequence that suggests a common mechanism for plasma membrane localization. Thus, it is possible that many of the plasma membrane isoforms are actually integrated into different isoform-specific regulatory complexes.
Subcellular targeting information presented here provides a starting point
for understanding the isoform-specific functions for nine of the 34 AtCPKs.
For AtCPKs 3 and 4, which may translocate between the cytosol and nucleus,
potential substrates include transcription factors. For AtCPKs 7, 8, 9, 16,
21, and 28, which are located at the plasma membrane, potential substrates
include ion pumps and channels (e.g., Li
et al., 1998
Plant Material and Transformation
All transgenic plants were generated using Arabidopsis ecotype Columbia.
The following CDPK-GFP fusion constructs were transformed into plants with
Agrobacterium tumefaciens (GV3101;
Koncz and Schell, 1986
CDPK cDNAs were amplified by PCR with the following gene-specific primers (the first and last codon of the cDNA is in bold, restriction sites are underlined): CPK1-5' (867A): 5'-CAGTACGTAAAACATGGGTAATACTTGTGTT3' and CPK1-3' (867B): 5'-CT-GGCGCGCCCTCTAGACCCATTTTCAC-3'; CPK3-5' (255A): 5'-AAGCTCGAGAC-AATGGGCCACAGACACAGCAAGTCCAAATCC-3' and CPK3-3' (255B): 5'-TCA-AACTAGTCCGCCCATTCTGCGTCGGTTTGGCAC-3'; CPK4-5' (256A): 5'-AAGC-TCGAGATGGAGAAACCAAACCCTAGA-3' and CPK4-3' (256B): 5'-TCAAA-CTAGTCCGCCCTTTGGTGAATCATCAGATTT-3'; CPK7-5' (257A): 5'-TTTGTCGACATGGGGAATTGTTGTGGCAAT-3' and CPK7–3' (257B): 5'-TCAAACTAGT-CCGCCGGTCTCGCCTTCTAATTGCAA-3'; CPK8-5' (258A): 5'-GAAGTCGACAT-GGGAAATTGTTGTGCGAGC-3' and CPK8-3' (258B): 5'-TCAAACTAGTC-CGCCATTTTCGCCTTCTAATTGCAA-3'; CPK9-5' (259A): 5'-CTTCTCGAG-GGCGCGCCGGGTATGGGAAATTGTTTTGCCAAG-3' and CPK9-3' (259B): 5'-CAAAACTAGTCCGCCGAACAGCCGAGGTTGTTGTTG-3'; CPK16-5' (735A): 5'-AAGCTCGAGATGGGTCTCTGTTTCTCCTCCGCCG-3' and CPK16-3' (735BR): 5'-CGGGTCTAGACCGCCACCGACCTTGCGAGAAATAAGATAACCA-3'; CPK21-5' (265A): 5'-GGAGTCGACCTCGAGGGTATGGGTTGCTTCAGCAGTA-AACACCGGAA-3' and CPK21-3' (266B): 5'-TTGGCCTAGGGGCGCGCCGCC-ATGGAATGGAAGCAGTT-3'; CPK28-5' (734A): 5'-AAGCTCGAGATGGGTGTC-TGTTTCTCCGCCATT-3' and CPK28-3' (734BR): 5'-CGGGTCTAGACCGCCACC-TCGAAGATTCCTGTGACCTGCAGG-3'.
All CDPK clones derived from PCR were sequenced to verify the absence of
mistakes. Fragments were cloned into the XhoI/SpeI site in
vector p35S-GFP-JFH1 (Hong et al.,
1999
AtCPK1-
The GFP constructs for Figure 3, B and D
to F, are based on Monsanto plant expression vector pMON10098
(Klee et al., 1991 GFP targeted to peroxisomes (seen in Fig. 3C) was accomplished by adding the plant peroxisome-targeting signal KSRM to the end of GFPrs. Primers oGFP12 (873A), 5'-GGACTAGTACGTAAAACATGAGTAAAGGAGAAGAA-3', and oGFP-KSRM (873B), 5'-CGGGATCCGCGGCCGCTCACATCCTGGATTTGTATAGTTCATCC-AT-3', were used to amplify GFPrs. The SpeI and BamHI sites at the ends of this PCR product were used to directionally clone the fragment into a modified pMON10098 vector. The resulting construct (pMG.px.r; PS#598) has GFPrs with the amino acids KSRM added to the C-terminal end, the 35S promoter driving the open reading frame, and an E9 terminator. The construct with two GFPs (used in Fig. 3, D–F) was prepared by first building AtCPK1 fused to GFPsg. AtCPK1 was amplified and cloned into pMG.rp1.sg. The resulting construct, pMG.AtCPK1.sg (PS#599), was combined with pMG.dpx.r (PS#597) to make a double GFP construct (pMG.dpx.r, is similar to pMG.px.r, except that it has an additional SfiI site for tandem cloning). The resulting construct pMG.px.AtCPK1 (PS#600) has both GFPrs-KSRM and AtCPK1-GFPsg expressed in tandem on the same plasmid.
Seeds from transgenic plants were selected on Murashige and Skoog plates (one-half-strength Murashige and Skoog salts and 0.5% [w/v] Suc, pH 5.7) containing 30 mg L–1 kanamycin. After 10 d, plants were transferred to Murashige and Skoog plates without antibiotics and were grown at 23°C with constant light. After 1 week, plants were used for imaging. Plants in Figure 4 were treated overnight with 10 µM dexamethasone before imaging. Images of CDPK-GFP fusion proteins in Figures 1, 3, B and C, and 4 were captured with a confocal laser scanning microscope (IX70 [Olympus, Tokyo]with Fluoview software). Excitation and emission filter peaks were 488 and 525 nm, respectively.
For covisualization of MitoTracker Red (Molecular Probes, Eugene, OR) and
AtCPK1-GFP, the image stack in Figure
3A was captured with a computational optical sectioning microscope
system with the 20x lens and the filters described by Gens et al.
(1996 Localization of AtCPK1-GFP and KSRM-GFP fusion proteins in Figure 3, D through F, was assessed with a wide-field fluorescence microscope fitted with controls and Slidebook software by Intelligent Imaging Innovations (Santa Monica, CA). Excitation filters were 405 and 490 nm, respectively, with a bandpass of 20 nm; the emission filter was 530 nm with a bandpass of 30 nm. Blur and fluorescence from the cytosol was reduced with the SlideBook no-neighbors algorithm. To check that our settings allowed a clear distinction between the two fluorophores of AtCPK1-GFPsg and GFPrs-KSRM, both constructs were expressed separately and imaged as described in Figure 3, D through F. In a second set of control experiments GFPsg and GFPrs were targeted to different organelles and were imaged in the same cell. No overlap of the fluorescent signals could be detected under the described conditions (data not shown).
Two-week-old transgenic Arabidopsis plants were further cultured in liquid Murashige and Skoog medium (one-half-strength Murashige and Skoog salts, 2% [w/v] Suc, and vitamin solution [Sigma, St. Louis], pH 5.7) at 22°C for 3 weeks in the dark on a rotary shaker. Roots were drained and frozen in liquid nitrogen. Samples were ground in a mortar and mixed with homogenization buffer (100 mM Tris-HCl, pH 7.5, 300 mM Suc, 10 mM EDTA, and 2 mM EGTA). To remove cell debris, the slurry was first passed through cheesecloth and then centrifuged for 15 min at 2,000g. All microsomal membranes were recovered from the supernatant by centrifugation at 100,000g for 1 h. The microsomal membrane pellet was resuspended in resuspension buffer (10 mM Tris-HCl, pH 7.5, 1 mM EDTA, and 1 mM EGTA). Membrane and soluble protein fractions were stored at –80°C. All buffers contained proteinase inhibitor cocktail (Roche Diagnostics, Mannheim, Germany).
Bradford assays were used to determine protein quantities (Bio-Rad Laboratories, Hercules, CA). Five to 10 µg of each sample was precipitated with 10% (w/v) trichloroacetic acid, resuspended in loading buffer, and separated on 10% or 14% (w/v) SDS-polyacrylamide gels. Proteins were then transferred to nitrocellulose membranes (Schleicher & Schuell, Keene, NH). Membranes were blocked overnight in Tris-buffered saline plus Tween 20 (10 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.1% [w/v] Tween 20 containing 5% [w/v] dry milk). The membranes were then probed with anti-GFP antibody (1:500 dilution; BD Biosciences Clontech, Palo Alto, CA) for 1 h at room temperature. After washing three times for 10 min in Tris-buffered saline plus Tween 20, membranes were probed with goat anti-rabbit antibody conjugated to horseradish peroxidase (1:5,000 dilution). Signals were detected using the SuperSignal West Femto chemiluminescent substrate kit (Pierce, Rockford, IL).
We thank Dr. Yan Wu (Torrey Mesa Research Institute, San Diego) for assistance with confocal microscopy. Received January 3, 2003; returned for revision February 5, 2003; accepted April 21, 2003.
Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.103.020008.
1 This work was supported in part by the Department of Energy (grant no.
DE–FG03–94ER20152 to J.F.H.), by the National Science Foundation
(grant nos. MCB011476 and IBN-9416038 to J.F.H.), by the Human Frontiers
Science Program (grant no. RG0268to J.F.H.), by Syngenta (to J.F.H.), by the
National Aeronautics and Space Administration/National Science Foundation
Joint Program in Plant Biology (grant no. NAGW 3046 to B.G.P.), and by the
National Science Foundation (grant no. MCB 9973770 to A.C.H.).
2 These authors contributed equally to the paper. * Corresponding author; e-mail Harper{at}Scripps.edu; fax 858–784–9840.
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ABSTRACT
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M).
z only occurs if the proteins are myristoylated and
palmitoylated (Alland et al.,
1994
