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Eukaryotic Cell, December 2008, p. 2100-2112, Vol. 7, No. 12
1535-9778/08/$08.00+0     doi:10.1128/EC.00118-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

C2 Domain Protein MIN1 Promotes Eyespot Organization in Chlamydomonas reinhardtii{triangledown} ,{dagger}

Telsa M. Mittelmeier,1* Peter Berthold,2 Avihai Danon,3 Mary Rose Lamb,4 Alexander Levitan,2 Michael E. Rice,1 and Carol L. Dieckmann1

Department of Biochemistry and Molecular Biophysics, University of Arizona, Tucson, Arizona 85721,1 Institute of Enzymology, Hungarian Academy of Science, 1113 Budapest, Hungary,2 Department of Plant Sciences, Weizmann Institute of Science, Rehovot 76100, Israel,3 Department of Biology, University of Puget Sound, Tacoma, Washington 984164

Received 2 April 2008/ Accepted 3 October 2008


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ABSTRACT
 
Assembly and asymmetric localization of the photosensory eyespot in the biflagellate, unicellular green alga Chlamydomonas reinhardtii requires coordinated organization of photoreceptors in the plasma membrane and pigment granule/thylakoid membrane layers in the chloroplast. min1 (mini-eyed) mutant cells contain abnormally small, disorganized eyespots in which the chloroplast envelope and plasma membrane are no longer apposed. The MIN1 gene, identified here by phenotypic rescue, encodes a protein with an N-terminal C2 domain and a C-terminal LysM domain separated by a transmembrane sequence. This novel domain architecture led to the hypothesis that MIN1 is in the plasma membrane or the chloroplast envelope, where membrane association of the C2 domain promotes proper eyespot organization. Mutation of conserved C2 domain loop residues disrupted association of the MIN1 C2 domain with the chloroplast envelope in moss cells but did not abolish eyespot assembly in Chlamydomonas. In min1 null cells, channelrhodopsin-1 (ChR1) photoreceptor levels were reduced, indicating a role for MIN1 in ChR1 expression and/or stability. However, ChR1 localization was only minimally disturbed during photoautotrophic growth of min1 cells, conditions under which the pigment granule layers are disorganized. The data are consistent with the hypothesis that neither MIN1 nor proper organization of the plastidic components of the eyespot is essential for localization of ChR1.


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INTRODUCTION
 
A variety of organelles and multicomponent structures are assembled and maintained within eukaryotic cells. These structures occupy defined, often asymmetric, intracellular locations and must be correctly partitioned or reassembled at every cell division. Elucidation of the mechanisms that govern the assembly and localization of complex structures is critical to our understanding of how basic cellular processes are regulated, how they may have evolved, and the factors driving their ongoing diversification.

Assembly and asymmetric localization of the photosensory eyespot in cells of the biflagellate, unicellular, photosynthetic green alga Chlamydomonas reinhardtii provide useful models for genetic, molecular, and microscopic analyses of organelle biogenesis (11, 14, 23). Chlamydomonas is phototactic, using two anterior flagella to swim toward or away from a light source to locations where light intensity is optimal for photosynthesis but minimally damaging to the photosynthetic membranes (82). The eyespot (Fig. 1A, wild type) is a light-sensing structure positioned near the equator of the cell at an asymmetric location relative to the flagella (25). Stimulation of the rhodopsin family photoreceptors in the eyespot activates a Ca2+-dependent signal transduction pathway(s) that affects flagellar movement and the swimming behavior of the cell (24, 58, 69). Asymmetric localization of the eyespot is required for the transmission of information about the direction of the light source (82).


Figure 1
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  		FIG. 1. Eyespots in wild-type and min1 mutant cells. (A) As previously described (33), a light micrograph of a min1 mutant cell (strain 12-12) reveals a miniature, equatorially localized eyespot (arrow). (B) Diagram of a wild-type eyespot showing layers of carotenoid pigment granules (dark gray circles) and thylakoid membrane (TM) immediately apposed to the inner and outer membranes of the chloroplast envelope (CE) (arrows) and the plasma membrane (PM). The eyespot photoreceptors (light gray ovals) are presumably in the plasma membrane.

The sensitivity of the eyespot to light is a result of the properties and organization of the photoreceptors and other eyespot components (Fig. 1B) (18, 30). In the eyespot, the plasma membrane is apposed to the chloroplast envelope and several underlying (plastid-localized) layers of carotenoid-filled granules subtended by thylakoid membranes (44). The pigmented granules give the eyespot its distinctive orange-red color when Chlamydomonas is viewed with a light microscope. The regularly spaced granule/thylakoid membrane layers function as a biological quarter-wave plate, reflecting orthogonal light back toward the rhodopsin photoreceptors in the plasma membrane (alternatively termed ChR1 and ChR2 [51, 52], CSRA and CSRB [74], and Acop1 and Acop2 [78]) and absorbing light from other directions. The defined location of the eyespot within the cell and the layered organization of the plastid and plasma membranes in the eyespot are directly observable examples of coordination that has evolved from association of a historical host with a cyanobacterial endosymbiont (15).

Here we describe the Chlamydomonas MIN1 gene, which is required for proper assembly of the eyespot. MIN1 was identified in a screen for mutant strains that were not phototactic and had missing or abnormal eyespots (33). min1 mutant cells have miniature eyespots located near the equator of the cell (Fig. 1A, min1). In cells grown photoautotrophically (in the light and without a source of reduced carbon, such as acetate), the carotenoid granules of min1 eyespots are disorganized and the chloroplast envelope is no longer apposed to the plasma membrane. Characterization of the MIN1 cDNA predicts a 322-residue protein with a novel domain organization that includes an N-terminal C2 (phospholipid-binding) domain and a C-terminal LysM (peptidoglycan-binding) domain. Similarities between the domains in MIN1 and those in membrane-associated proteins and localization of the C2 domain to the chloroplast envelope in moss cells support the hypothesis that MIN1 promotes membrane apposition in the eyespot via direct interaction with the chloroplast envelope and/or plasma membrane. Analyses of channelrhodopsin-1 (ChR1) photoreceptor levels and localization in min1 mutants suggest that MIN1 also promotes expression and/or stabilization of ChR1. Finally, the data are consistent with a model in which ChR1 localization is not wholly dependent upon proper organization of the pigment granule layers of the eyespot.


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MATERIALS AND METHODS
 
Strains and media. Chlamydomonas reinhardtii min1 mutant strain 12-12 (137c min1 mt+) (33) was obtained by UV mutagenesis of strain 137c (21), followed by screening for mutants that were not phototactic (ptx–). Strain 12-12 arg2+ (min1 arg2 mt+) was a spore from a cross of strain 12-12 to strain arg2– (137c arg2 mt–) (21).

Chlamydomonas cultures were maintained on solid TAP medium (21) or on TAP medium plus 0.2 mg/ml arginine. For phototaxis assays, Western blot analyses of total cellular protein, or microscopy, liquid cultures were grown photoautotrophically in modified Sager and Granick medium I with Hutner's trace elements (M medium) (22) or mixotrophically in the same medium containing 0.1% sodium acetate (R medium). For isolation of genomic DNA, liquid cultures were grown in R medium. For transformation of strain 12-12 arg2+, liquid cultures were grown in R medium limited for NH4NO3 (0.125 mM) and supplemented with 0.2 mg/ml arginine (RNA medium). All cultures were grown at 25°C under continuous light.

Phototaxis assays. Following overnight growth in liquid M medium, a simple assay in which phototactic cells in a test tube swim toward a lighted slit at the bottom of an otherwise dark box (33) was used to determine whether cells were phototactic (ptx+) or not (ptx–).

Identification of the MIN1 gene. The min1 mutant strain H6-2 (137c min1::ARG7 mt+) was isolated from an ARG7 (arginosuccinate lyase) insertion library (2, 66) and crossed to strain arg7– (137c arg7 mt–) (21). Phenotypic analysis of 20 tetrads and 21 random spores showed that the ARG7 insertion in H6-2 was linked to min1 (data not shown). Plasmid rescue was used to recover genomic sequence neighboring the ARG7 insertion in H6-2 (66, 79). Probes derived from the recovered sequence identified a restriction fragment length polymorphism that segregated with the min1 phenotype (data not shown), confirming linkage to min1.

Oligonucleotides 5B-22 and 5B-783 (see Table S1 in the supplemental material), derived from the recovered genomic sequence, identified cosmid H5b from a pARG7.8cos cosmid library (63). Following transformation of strain 12-12 arg2+ with linearized H5b, 7% of the Arg+ transformants (4/60) were phototactic (ptx+) and had normal eyespots when viewed by light microscopy (data not shown). A 5.0-kb BamHI-HpaI fragment from H5b was ligated to BamHI- and EcoRV-digested pARG7.8 to construct pARG7.8-MIN1BH. Following transformation of strain 12-12 arg2+ with pARG7.8-MIN1BH (pMIN1 in Table 1), 12.5% of Arg+ transformants (25/200) were phototactic and had normal eyespots. The sequence of both strands of the 5.0-kb insert in pMIN1 was determined by progressive design of oligonucleotide primers (see Table S1 in the supplemental material). Comparison of the 5.0-kb sequence to version 3.0 of the Chlamydomonas genome sequence (48) at the DOE Joint Genome Institute (JGI) yielded a match on scaffold 11 (model 11000167).


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  		TABLE 1. MIN1 constructs and phototaxis rescuea

Identification of the min1 mutation in strain 12-12. Genomic DNA isolated from the min1 mutant strain 12-12 was used as the template in PCRs using overlapping oligonucleotide primer pairs (see Table S1 in the supplemental material), and the PCR products were ligated to the pGEM-T Easy plasmid (Promega, Madison WI) and sequenced. Repeated amplification with primer pair A7 plus B7 verified the G-to-T transversion at codon E61 of the predicted MIN1 protein, which changed a GAG codon to a premature TAG termination codon.

Characterization of the MIN1 cDNA. The 5.0-kb sequence containing MIN1 was searched for probable coding regions using the GeneMark gene prediction algorithm (39), and oligonucleotide primers (see Table S1 in the supplemental material) were used in PCR amplification of MIN1 cDNAs from a Chlamydomonas cDNA library made from mRNA isolated from synchronized cells just after cell division (G. Pazour and G. Witman, personal communication). Amplification required two 25-cycle reactions; 1/10 of the first reaction product was used as the template in the second reaction. The PCR products were ligated to the pGEM-T Easy plasmid and sequenced. The 5' end of the cDNA was defined by primers B7.5 (100 nucleotides [nt] 5' of the ATG), which consistently yielded a PCR product, and B7.3 (144 nt 5' of the ATG), which did not. The 3' end of the cDNA extends beyond primer A2, approximately 1 kb 3' of the termination codon. The deduced exon/intron structure of the MIN1 gene is shown in Fig. 2.


Figure 2
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  		FIG. 2. The Chlamydomonas MIN1 gene and the RbcS2-MIN1i7 and HA-tagged constructs. (A) Organization of the 5.0-kb genomic sequence in pMIN1 (see Materials and Methods and Table 1). White boxes are UTRs. TAG 12-12 indicates the position of the nonsense mutation at codon E61 (in exon 3) in the min1 mutant strain 12-12. (B) The RbcS2-MIN1i7 translational fusion in pR-MIN1 (see Materials and Methods and Table 1). The RbcS2 promoter and intron 1 were ligated in frame to MIN1 coding sequence containing only intron 7. In the HA-tagged constructs (Table 1), sequence encoding the triple HA epitope was ligated in frame at codon 249, just 5' of sequence encoding the predicted transmembrane domain. (C) Light micrographs of a wild-type cell and of min1 cells transformed with pMIN1-HA, pR-MIN1, or pR-MIN1-HA (see Materials and Methods and Table 1). The cultures were grown in M medium, and the single cells shown were typical of the majority of cells in each culture. (D) Western blot of total cellular protein isolated from M medium-grown cultures of untransformed strain 12-12 (min1) or of transformant strains containing the indicated constructs. The blot was probed with anti-HA (MIN-HA) (clone 12CA5; Sigma, St. Louis, MO), followed by antitubulin (clone B-5-1-2; Sigma). Shorter (top) and longer (middle) exposures of the blot probed with anti-HA are shown. pR-MIN1-HA strains 68 and 70 were obtained following three rounds of enrichment for ptx+ cells.

DNA sequencing. Automated DNA sequencing was performed at the DNA Sequencing Facility of the Arizona Research Labs at the University of Arizona (Tucson, AZ).

Plasmid construction. Plasmid pR-MIN1 (Table 1 and Fig. 2) was constructed by PCR amplification of MIN1 cDNA sequence from the predicted start codon (primer MIN1-Nde-start [see Table S1 in the supplemental material]) to the BsrG1 site just 5' of intron 7 (primer MIN1-BsrG1A) and amplification of the RbcS2 promoter plus intron 1 sequence from plasmid NE-537 (GenBank accession AY710294 [GenBank] ) (68) using primers BamH1-RbcS2 and EcoRV-RbcS2A. The amplification products were ligated to create an in-frame fusion of MIN1 coding sequence to the RbcS2 sequence, and the fusion construct was used to replace the genomic BamHI-to-BsrG1 sequence in pMIN1. pMIN1-HA and pR-MIN1-HA were constructed by ligation of a PCR-amplified triple-hemagglutinin (HA) tag sequence to MscI-digested pMIN1 or pR-MIN1 (the MscI site is at codons 248 to 250, just 5' of sequence encoding the transmembrane domain). Plasmids containing mutations in MIN1 ({Delta}3'ATG, D19A,D77A, K37A,K42A, and T38A) were constructed as follows: MIN1 sequence was amplified by PCR using oligonucleotide primer pairs (see Table S1 in the supplemental material) that replaced wild-type sequence with sequence that altered a single codon and created a unique restriction site, which was used to assemble the fragments, and wild-type sequence in either pMIN1 (for pMIN1-{Delta}3'ATG) or pR-MIN1 (for pR-D19A,D77A, pR-K37A,K42A, or pR-T38A) was replaced with the mutant MIN1 sequence.

Plasmids {Delta}LMTM-YFP and {Delta}LMTMda (used for transient transfection of moss cells) were constructed by in-frame fusion of PCR-amplified (oligonucleotide primers are listed in Table S1 in the supplemental material) MIN1 cDNA sequence encoding residues 1 through 180 to PCR-amplified yellow fluorescent protein (YFP). The fusion was ligated to a moss expression vector containing the actin promoter and transcription termination sequences (36).

Chlamydomonas transformation. Chlamydomonas strain 12-12 arg2+ was transformed using silicon carbide whiskers (13) as previously described (66) with the following modification: following growth in liquid RNA medium to approximately 2 x 106 cells/ml, the cells were harvested by centrifugation, resuspended in 200 ml of low-nitrogen medium, and grown overnight at 25°C under continuous light.

Fluorescence microscopy of moss cells. Physcomitrella patens B. S. G. was grown on solid minimal NH4 medium at 25°C on a 16-hour/8-hour light/dark cycle. For transformation of P. patens (36), protoplasts were isolated from 5- to 6-day-old protonemal cultures and 10 µg of plasmid DNA was added to 300 µl of a protoplast suspension. Following gentle mixing, 300 µl of a solution containing 40% polyethylene glycol, 0.1 M CaNO3, 0.38 M mannitol, and 10 mM Tris-HCl (pH 8.0) was added, and the suspension was incubated with occasional mixing for 5 min at 45°C and for 10 min at room temperature. The protoplast suspension was diluted to a final volume of 7.1 ml with liquid NH4 medium supplemented with 6.8% mannitol and incubated for 16 h in the dark. Fluorescence images were taken 24 h after transformation using an Olympus Fluoview FV500 laser confocal microscope. YFP, cyan fluorescent protein (CFP), and chlorophyll autofluorescence images were obtained using excitation/collection wavelengths of 514/560 to 615 nm (YFP), 405 to 445/460 to 500 nm (CFP), and 630 nm (chlorophyll).

Light microscopy. Inocula from fresh cultures on solid medium were transferred to 2 ml of liquid M medium and grown at 25°C for 2 days under continuous light. The cells were viewed with a Leica DMRXA microscope using a Leica PL APO 100x, 1.4-numerical-aperture oil immersion objective with a 1.6x optivar (1 pixel = 0.039 µm) and bright-field optics. Images were captured with a QImaging (Burnaby, British Columbia, Canada) Retiga EX-cooled charge-coupled device camera driven by Universal Imaging (Downingtown, PA) MetaMorph v.6.1.2 software.

To determine eyespot area (in pixels), the MetaMorph (Universal Imaging, Downingtown, PA) "threshold image" function was used to select pixels representing the eyespot, and the "morphometric analysis" of "single objects" function was used to count the number of pixels selected. The eyespot was in the plane of focus in all images. Additionally, the entire circumference of the eyespot was visible in all images used for this analysis.

Western blotting. Inocula from fresh cultures on solid medium were transferred to 2 ml of liquid M or R medium and grown overnight at 25°C under continuous light. The cells were harvested at 20,800 x g for 10 min, resuspended in 100 µl to 200 µl of 4x Laemmli buffer (250 mM Tris-Cl [pH 6.8], 40% glycerol, 20% β-mercaptoethanol, 8% sodium dodecyl sulfate, 0.024% bromophenol blue) (32) with protease inhibitors (5 µg/ml aprotinin, 5 µg/ml leupeptin, 1 µg/ml pepstatin A, and 1.0 mM phenylmethylsulfonyl fluoride), and heated at 100°C for 5 min. Thirty microliters of each sample was electrophoresed through 10% polyacrylamide-sodium dodecyl sulfate gels and transferred to BioTrace polyvinylidene difluoride membranes (Pall Corp., Ann Arbor, MI) using standard techniques. The blots were blocked in 5% nonfat dry milk (NFDM) in TBS-T (10 mM Tris-Cl, 150 mM NaCl, 0.5% Tween 20) for 1 h at room temperature; probed overnight at 4°C with either rabbit polyclonal anti-ChR1 (1:5,000) (5), mouse antitubulin (clone B-5-1-2 at 1:10,000; Sigma, St. Louis MO), or mouse anti-HA (clone 12CA5; 1:1,000, Sigma) in 1% NFDM in TBS-T; washed in TBS-T; and probed with a 1:10,000 dilution of either goat anti-rabbit-horseradish peroxidase (Pierce, Rockford IL) or goat anti-mouse-horseradish peroxidase in 1% NFDM in TBS-T. Following several washes in TBS-T, the blots were incubated in SuperSignal substrate (Pierce) for 1 min and exposed to ECL Hyperfilm (Amersham Biosciences, Piscataway, NJ). The "integrated density" function of ImageJ software was used to measure the intensity of individual bands in digital images of Western blots obtained using a PaperPort scanner. The anti-ChR1 signal obtained from each sample was normalized to the antitubulin signal in the same sample.

Immunofluorescence. Inocula from fresh cultures on solid medium were transferred to 2 ml of liquid M or R medium and grown overnight at 25°C under continuous light. Cells were harvested from 0.5 ml of culture at 2,700 x g for 10 min, resuspended in Chlamydomonas autolysin prepared from strains 4A+ and 1B– (kind gifts of Patrice Hamel, Ohio State University, Columbus, OH), and incubated for 1 h at room temperature. The cells were harvested, resuspended in phosphate-buffered saline (PBS), spotted onto 10-well poly-L-lysine-coated slides, allowed to settle for 10 min at room temperature, and then dipped into –20°C methanol for 10 s. After a brief drying period, the cells were incubated in block buffer (1x PBS, 0.1% Tween 20, 1% bovine serum albumin) for 1 h at room temperature and incubated with rabbit anti-ChR1 (a gift of P. Berthold and P. Hegemann), diluted 1:50 in block buffer, overnight at 4°C. The cells were then washed four times for 10 min each in wash buffer (block buffer without bovine serum albumin), incubated in 1:400 donkey anti-rabbit Alexa 488 (Molecular Probes, Eugene, OR) for 2 h at room temperature, washed three times for 10 min each in wash buffer and once for 10 min in PBS, and coverslipped with VectaShield hard-set mounting medium (Vector Laboratories, Burlingame, CA). Alexa 488 fluorescence was viewed with a Leica DMRXA microscope using a Leica PL APO 100x, 1.4-numerical-aperture oil immersion objective with a 1.6x optivar (1 pixel = 0.039 µm) and a Chroma 71001A filter set (Chroma Technology Corp., Rockingham, VT). One- or 2-second exposures were captured using a QImaging (Burnaby, British Columbia, Canada) Retiga EX cooled charge-coupled device camera driven by Universal Imaging (Downingtown, PA) MetaMorph v.6.1.2 software. The images shown are summed maxima of Z-series (each Z-series contained 6 to 10 images, captured at 0.5-µm intervals) that were adjusted for brightness and cropped.

Figure preparation. Figures were produced using Microsoft Word, Adobe Photoshop, Adobe Illustrator, or a combination of these programs. Micrograph or immunoblot images were minimally adjusted for grayscale levels or brightness and contrast, cropped, and reduced from the original size.

Nucleotide sequence accession number. The sequence of the 5.0-kb insert in pMIN1 is available from GenBank/EMBL/DDBJ under accession number AY45207.


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RESULTS
 
The MIN1 gene. MIN1 was originally identified by genetic analysis of a phototaxis-deficient strain with miniature, disorganized eyespots in which the plasma membrane and chloroplast envelope are not apposed (Fig. 1) (33). Measurement of eyespot area (in pixels; see Materials and Methods) in digital light micrographs indicated that min1 eyespots (39 ± 11 pixels, n = 33) cover approximately 25% of the area covered by wild-type eyespots (148 ± 35 pixels, n = 46). To identify MIN1, Chlamydomonas genomic sequence neighboring the ARG7 insertion in strain H6-2 (137c arg2 min1::ARG7) was isolated and used to identify a 5.0-kb BamHI-HpaI fragment of the genome that, following ligation to the pARG7.8 vector, restored phototaxis (ptx+) in 17% of the Arg+ transformants (Table 1). Light microscopy confirmed that the ptx+ transformants had eyespots of normal size (data not shown). The sequence of the MIN1-containing fragment (GenBank accession AY452057 [GenBank] ) matched sequence on scaffold 11 of the DOE Joint Genome Institute Chlamydomonas reinhardtii genome sequence (JGI version 3.0, http://genome.jgi-psf.org/Chlre3/Chlre3.home.html) (48) approximately 913,000 bp from MLT1 (multieyed) (33; T. M. Mittelmeier and C. L. Dieckmann, unpublished data) and 958,000 bp from EYE2 (eyeless) (66). The 7% or 9% recombination observed between MIN1 and the MLT1 or EYE2 locus, respectively (33), is consistent with the recombination rate of 100 kb per cM estimated for the Chlamydomonas genome (48).

To identify the MIN1 coding sequence, oligonucleotide primers (see Table S1 in the supplemental material) based on open reading frame (ORF) predictions made by the GeneMark algorithm (39) were used to amplify MIN1 cDNAs by PCR (see Materials and Methods). The MIN1 cDNA is approximately 2.1 kb, comprising eight exons and seven introns (Fig. 2A), and includes a 322-codon ORF. In the original min1 mutant strain 12-12, the 322-codon ORF is interrupted by a termination codon resulting from a GAG-to-TAG transversion at codon 61 (data not shown), consistent with the prediction that this ORF encodes MIN1. The 5' splice junctions have the sequence R/GT (R = A or G), while the 3' splice junctions have the sequence RCAG/N, which conform to proposed consensus splice site sequences (63). The approximate 5' and 3' ends of the mRNA were defined by oligonucleotide primer pairs that did or did not yield amplified cDNAs (see Materials and Methods). The 5' untranslated region (UTR) of the cDNA is less than 150 nt long, while the 3' UTR is over 1 kb with the consensus polyadenylation sequence TGTAA (47, 73) 1,122 nt downstream of the stop codon. The 3' UTR also contains two short overlapping ORFs (84 and 74 codons), but the rate of phenotypic rescue was unaffected by an ATG-to-CTG mutation in the start codon of the longer ORF (Table 1, pMIN1{Delta}3'ATG), indicating that this ORF, if expressed, is not required for eyespot assembly or function. A construct in which the MIN1 coding sequence, containing only the final intron, was fused to the RbcS2 promoter and first intron (Fig. 2B) (20, 40) consistently yielded a rescue rate of nearly 50% (Table 1, pR-MIN1). Thus, the genomic MIN1 promoter and 5'UTR are not required for functional expression of the MIN1 protein.

The size of a MIN1-HA protein is consistent with the predicted MIN1 ORF. In an attempt to localize the MIN1 protein in Chlamydomonas, epitope-tagged MIN1 constructs were assessed for phenotypic rescue. Constructs expressing the MIN1-coding sequence fused to C-terminal tags did not rescue the min1 phenotype (data not shown). Integration of the triple-HA tag (17) just N terminal of the predicted MIN1 transmembrane sequence in the context of either the genomic MIN1 clone (pMIN1-HA) or the RbcS2-MINi7 fusion construct (pR-MIN1-HA) yielded ptx+ transformants (Table 1). However, the rate of rescue by pMIN1-HA was relatively low, pMIN1-HA transformant cultures displayed weak phototaxis and had miniature eyespots following overnight growth (Fig. 2C), and both pMIN1-HA and pR-MIN1-HA transformants eventually lost the ptx+ phenotype. These data suggest that the HA tag negatively affected the phenotypic expression of the MIN1 gene, either by reducing expression of the transgene, perhaps by increasing silencing (8), and/or by compromising function of the MIN1 protein.

To allow further analysis of the HA-tagged MIN1 protein, two pR-MIN1-HA transformants were subjected to three rounds of selection for ptx+ cells following sequential phototaxis assays (see Materials and Methods). The enrichment yielded the ptx+-stable strains pR-MIN1-HA-68 and -70 (Fig. 2C); the enrichment may have favored cells in which the transgene was not silenced. The anti-HA monoclonal antibody 12CA5 detected a 35-kDa protein on Western blots of total cellular protein from strains pR-MIN1-HA-68 and -70 and from a pMIN1-HA transformant following overexposure of the blot. The 35-kDa protein was close in size to the 37.7 kDa predicted for MIN1-HA (34.2 kDa for MIN1 plus 3.5 kDa for the triple-HA tag) and was not detected in transformants containing the untagged construct, confirming its identity as the MIN1-HA fusion protein. These data are consistent with the conclusion that the 322-codon ORF within the MIN1 cDNA encodes the MIN1 protein.

While the MIN1-HA protein was detectable by Western blotting in the enriched strains, repeated attempts to localize the MIN1-HA protein in Chlamydomonas by immunofluorescence using several anti-HA monoclonal antibodies (12CA5 from Boehringer Mannheim, 3F10 from Roche, or HA.C5 from AbCam) were not successful (data not shown). As the usefulness of the HA-tagged construct was limited, alternative approaches will be necessary to characterize the expression and localization of the MIN1 protein.

The MIN1 protein contains three conserved domains. Queries of the NCBI databases using blastp (1, 41) identified two conserved domains within the predicted MIN1 protein (Fig. 3A). The N-terminal 121 residues had significant similarity to Ca2+/phospholipid-binding C2 domains SMART00239.7 (35, 72) and pfam 00168 (4), while the C-terminal 46 residues were similar to LysM domain sequences (pfam 01476) that bind peptidoglycan components of the bacterial cell wall (3, 6, 77). The secondary-structure prediction program TMHMM (trans-membrane helix prediction using the hidden Markov model) (31) identified residues 251 through 270 as a probably membrane-spanning {alpha}-helix. Residues 122 through 234 of MIN1 comprise an alanine-rich region (26% alanine), a common feature of Chlamydomonas proteins due to the high GC content of the genome (48). To date, the domain architecture of MIN1 is unique; no other proteins containing both a C2 domain and a LysM domain were identified by searches of pfam or the Conserved Domain Database (43).


Figure 3
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  		FIG. 3. The MIN1 protein contains C2, transmembrane, and LysM domains. (A) The domain structure of the MIN1 protein as determined by BLAST alignment to C2 (SMART00239.7) and LysM (pfam 01476.8) domain family members. The solid black region, from residue 251 to 270 of MIN1, is predicted to form a membrane-spanning {alpha}-helix. (B and C) ClustalW alignments. Asterisks denote positions at which all of the sequences have identical residues. Dots denote conservation of residues in ≥50% of the sequences. (B) ClustalW alignment of the MIN1 transmembrane domain with predicted membrane-spanning sequences identified using blastp. The GenBank accession number follows each sequence: Nocardiodes sp. strain JS614 sulfate transporter, Natronomonas pharaonis hypothetical protein NP2264A, Nocardioides sp. strain JS614 cobalmin-5-phosphate synthase, Dinoroseobacter shibae phosphate transporter, Chlamydomonas reinhardtii MIN1 protein, and Stigmatella aurantiaca Na+/H+ antiporter NhaA. (C) ClustalW alignment of the MIN1 LysM domain with LysM domain sequences identified using blastp. The GenBank accession number follows each sequence: Chlamydomonas reinhardtii MIN1, Moorella thermoacetica predicted protein, Ostreococcus tauri predicted protein Ot12g0040, Roseovarius nubinhibens LysM/phospholipid-binding domain protein, and Ralstonia solanacearum hypothetical protein RSc2148.

ClustalW (80) alignment of the MIN1 predicted transmembrane sequence or LysM domain sequence to protein sequences identified by blastp searches of GenBank are shown in Fig. 3B and C. The transmembrane sequence was similar to sequences in eubacterial hypothetical plasma membrane proteins, including probable sulfate and phosphate transporters from Nocardioides and Dinoroseobacter and an Na+/H+ antiporter from Stigmatella. The MIN1 LysM domain was most similar to the LysM domain in a hypothetical protein from the photosynthetic (but eyespot-less) alga Ostreococcus tauri (50% identity), the smallest known autotrophic eukaryote (59), and to those in hypothetical proteins from the eubacteria Moorella, Roseovarius, and Ralstonia (Fig. 3C). The aligned Ostreococcus and Moorella proteins each contain a predicted transmembrane helix, while the proteins from Roseovarius and Ralstonia contain BON (bacterial OsmY and nodulation; pfam04972) domains, which are hypothesized to bind phospholipids (83). Of interest is the observation that the LysM domain sequences most similar to that of MIN1 are in proteins predicted to be associated with membranes.

The MIN1 N terminus is a C2 domain capable of membrane association. C2 domains are found in proteins from a wide variety of eukaryotic organisms (and in the alpha-toxin of Clostridium perfringens) (54) that are either located in, or transiently associated with, cellular membranes (9, 65). C2 domains bind phospholipids, often in a Ca2+-dependent manner (34, 53). The MIN1 C2 domain sequence is most similar to hypothetical proteins from the red flour beetle (30% identity with the MIN1 C2 domain), the chicken (23%), the nematode Caenorhabditis (26%), the protozoan Leishmania (26%), the parasitic protozoan Trypanosoma (24%), and plants (Fig. 4A). With the exception of the Leishmania and Trypanosoma proteins, each of the aligned proteins was predicted by TMHMM to contain a transmembrane sequence. The Arabidopsis (25% identity) and rice (Oryza sativa, 25% and 26% identity) proteins also contain GRAM domains (named after the glucosyltransferases, Rab-like GTPase activators, and myotubularins that contain the domain), which were originally identified in eukaryotic proteins that function in membrane-associated processes (12). As was the case with the LysM domain, the MIN1 C2 domain sequence is most similar to that of C2 domains in proteins that are predicted to be associated with membranes.


Figure 4
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  		FIG. 4. The MIN1 N terminus is a predicted C2 domain. (A) ClustalW alignment of the MIN1 C2 domain with similar C2 domain sequences identified using blastp. Asterisks denote positions at which all of the sequences have identical residues. Dots denote conservation of residues in ≥50% of the sequences. MIN1 residues D19, D77, K37, K42, and T38 are indicated by large asterisks. The GenBank accession number of each aligned protein follows the sequence: Tribolium castaneum (red flour beetle) predicted protein, Gallus gallus (chicken) predicted protein, Caenorhabditis elegans hypothetical protein T12A2.15a, Leishmania infantum hypothetical protein LinJ31.0710, Trypanosoma cruzi hypothetical protein, Arabidopsis thaliana C2/GRAM domain protein At1G03370, Oryza sativa C2/GRAM domain protein (rice 08g0492400), and Oryza sativa C2/GRAM protein (rice 02g0199800). (B) ClustalW alignment of the MIN1 C2 domain with Brookhaven Protein DataBank sequences 1wfj (Arabidopsis C2 domain-containing protein from a putative elicitor-responsive gene) and 1rlw [PDB] (50) (C2 domain from Homo sapiens phospholipase A2) (60). Structurally determined (PDB sequences) and predicted (MIN1) β-sheet residues are italicized. Asterisks above the sequence indicate conserved residues D19, K37, T38, K42, and D77. (C) Ribbon diagram of the three-dimensional fold of the MIN1 C2 domain, predicted by the LOOPP algorithm (45, 81), based on the structure of the C2 domain in an Arabidopsis putative elicitor-responsive protein (PDB file 1wfj) (50). The diagram, produced using PyMOL (http://www.pymol.org) (10), illustrates the eight β-strands (1 through 8) of the C2 domain sandwich, the residues corresponding to the conserved loop aspartates in Ca2+-dependent domains (D19, N24, Q71, G73, and D77), and conserved residue T38, discussed in the text.

C2 domain sequences form a sandwich of two β-sheets, each comprised of four β-strands arranged in one of two distinct topological folds (26). In topology I structures, such as those in synaptotagmins (16), the most N-terminal β-strand (β1) occupies the same position in space as the most C-terminal strand (β8) in topology II structures, such as that of phospholipase A2 (60). In many C2 domains, three loops on one end of the sandwich contain five conserved aspartate residues that coordinate Ca2+ and are important for phospholipid binding (42, 70). Other C2 domains associate with phospholipid membranes in a Ca2+-independent manner, and nonacidic residues substitute for one or more of the loop aspartates (38, 55, 75). In protein kinase C{alpha}, a cluster of lysine residues in strands β3 and β4 (of a topology I domain) are also important for membrane binding (67).

The 3D-PSSM (28) and LOOPP (45, 81) threading algorithms predicted that the three-dimensional structure of the MIN1 C2 domain is most similar to those of the topology II C2 domains in human phospholipase A2 (Brookhaven Protein Database [PDB] file 1rlw [PDB] [60]; 15% identity with the MIN1 C2 domain) and an Arabidopsis putative elicitor-responsive protein (PDB file 1wfj [50]; 18% identity). An alignment of the MIN1 sequence to the phospholipase A2 and Arabidopsis sequences and the predicted fold of the MIN1 C2 domain, based on the structure of the Arabidopsis protein, are shown in Fig. 4B and C. Two loop aspartate residues are conserved in MIN1 (D19 and D77, in loops β1-β2 and β5-β6), while the remaining three loop aspartates have been replaced with amino acids that are theoretically capable of Ca2+ coordination (N24, Q71, and G73) (61). Three lysine residues (K37, K42, and K51) are clustered on one side of the sandwich in β-strands 3 and 4. K37 is highly conserved (9/10 sequences) and K42 is somewhat conserved (5/10 sequences) in C2 domains with the greatest similarity to that of MIN1 (Fig. 4A). Finally, residue T38 of MIN1 (Fig. 4) corresponds to a highly conserved threonine residue within strand β3 (topology II domains) (62) that may be important for proper folding of the domain.

The MIN1 C2 domain associates with the chloroplast envelope in moss cells. To investigate the potential function of the MIN1 C2 domain, plasmid DNA encoding the N-terminal 180 residues of MIN1 fused in frame to YFP ({Delta}LMTM-YFP) was used to transiently transfect moss cells (Physcomitrella patens) (36), and the resulting pattern of YFP fluorescence was analyzed microscopically (Fig. 5A and B). Similar to an Arabidopsis outer chloroplast envelope protein (GenBank accession no. 18419973)-CFP fusion protein, YFP fluorescence was associated with the chloroplast envelope when YFP was fused to the MIN1 C2 domain (Fig. 5) but not when YFP was used alone (data not shown). Changing residues D19 and D77, which correspond to conserved Ca2+-binding residues (Fig. 4), to alanine abolished association of the MIN1 C2-YFP fusion protein with the chloroplast envelope (Fig. 5). These data support the hypothesis that the MIN1 N terminus is a C2 domain that is potentially membrane associated, and they suggest that conserved loop aspartates are involved in the interaction.


Figure 5
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  		FIG. 5. The MIN1 C2 domain is associated with the chloroplast envelope in moss cells. (A) Physcomitrella patens (moss) cells were transiently transfected with plasmid DNA encoding a MIN1 C2 domain-YFP fusion protein containing either wild-type sequence ({Delta}LMTM:YFP) or sequence encoding a C2 domain in which conserved aspartic acid residues D19 and D77 were changed to alanines ({Delta}LMTMda:YFP). Cells expressing the YFP constructs were analyzed using an Olympus Fluoview FV500 laser confocal microscope. Green indicates YFP fluorescence, and red indicates chlorophyll autofluorescence. (B) High-magnification view of cells transformed with the MIN C2 domain-YFP constructs and plasmid DNA encoding an Arabidopsis outer chloroplast envelope protein (GenBank accession number 18419973)-CFP fusion protein. Green indicates YFP fluorescence, red indicates chlorophyll autofluorescence, and blue indicates CFP fluorescence.

Conserved residues within the C2 domain are not essential for eyespot assembly. To determine whether the membrane-binding function of the MIN1 C2 domain observed in moss cells is required for eyespot assembly, the D19A and D77A mutations were introduced into the RbcS2-MIN1i7 fusion construct in pR-MIN1 (see Materials and Methods) and the phenotypic consequences of the changes were assessed in Chlamydomonas. The aspartate residue mutations had no effect on the rate of ptx+ rescue (Table 1, pR-D19A,D77A), and eyespots in the transformed cells had a size similar to that of eyespots in wild-type cells (Fig. 6). Likewise, changing conserved lysine residues K37 and K42 to alanines or changing residue T38 to alanine had little to no effect on phenotypic rescue (Table 1 and Fig. 6, pR-K37A,K42A and pR-T38A). These data suggest that in Chlamydomonas, any interaction between the MIN1 C2 domain and eyespot membranes either has atypical requirements or is not essential for eyespot assembly, possibly because the function is redundant with that of another eyespot protein.


Figure 6
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  		FIG. 6. Conserved residues in the MIN1 C2 domain are not essential for eyespot assembly. Light micrographs of a wild-type cell or of min1 cells transformed with pR-D19A,D77A or pR-T38A (see Materials and Methods and Table 1) are shown. The cultures were grown in M medium, and the single cells shown were typical of the majority of cells in each culture.

ChR1 photoreceptor levels are dependent on MIN1. To determine whether the loss of MIN1 affects levels or localization of ChR1, one of the major photoreceptors in the eyespot (5, 51, 74), polyclonal anti-ChR1 (5) was used to analyze ChR1 levels and localization in wild-type or min1 mutant cells. On Western blots of Chlamydomonas whole-cell extracts from cells grown photoautotrophically (in the light in acetate-free medium, conditions under which the min1 eyespot is disorganized), the level of ChR1 was reduced to 65% ± 15% in the min1 nonsense mutant (strain 12-12) compared to the wild type (Fig. 7A) (see Materials and Methods). ChR1 levels were unaffected by either the RbcS2-MIN1 fusion or the mutation of conserved residues within the MIN1 C2 domain. These data indicate that MIN1 promotes photoreceptor expression and/or stabilization and that the conserved residues analyzed are not essential for this function.


Figure 7
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  		FIG. 7. Photoreceptor levels are low in min1 mutant cells. (A) Western blot of total cellular protein from M medium-grown cultures of wild-type (wt) or min1 mutant (strain 12-12) cells or of min1 cells transformed with pR-MIN1, pR-D19A,D77A, pR-K37A,K42A, or pR-T38A (see Materials and Methods and Table 1). The blot was probed with a polyclonal antibody against the ChR1 photoreceptor (5), followed by antitubulin (clone B-5-1-2; Sigma, St. Louis, MO). (B) Immunofluorescence of M medium-grown wild-type cells or min1 mutant cells (strain 12-12) probed with anti-ChR1. In the photographs of wild-type cells, the arrows point to an eyespot-associated "stripe" of immunofluorescence regularly observed in both wild-type and min1 cells. In the photograph of min1 cells, the arrow points to a cell containing two roughly equatorial "spots" of anti-ChR1 signal. The signal at the basal bodies is most likely nonspecific binding of anti-ChR1 (5).

Immunofluorescence with anti-ChR1 supported and extended these data (Fig. 7B). As observed previously, ChR1 was localized to a single equatorial spot in wild-type cells (5). Anti-ChR1 staining of the basal bodies observed in both wild-type and min1 mutant cells was most likely due to nonspecific binding of the anti-ChR1 polyclonal antibody, as it was observed in cells in which ChR1 expression had been silenced (5). In the min1 nonsense mutant, ChR1 spots were significantly smaller than those in the wild type, and in approximately one-fourth of the cells the fluorescence was distributed among two or more spots. However, the anti-ChR1 staining remained localized near the equator and to a single side of the cells. Given the disorganization of the pigment granule layers in min1 cells grown without acetate (33), these observations suggest that approximate localization of ChR1 does not require proper organization of the plastid components of the eyespot.

In every staining of either wild-type or min1 cells, we also observed cells with a longitudinal "stripe" of anti-ChR1 signal extending from the anterior end of the cell toward, or just next to, the eyespot (Fig. 7B) and sometimes beyond. The percentage of cells in which this stripe was observed varied between experiments, and additional data are required to determine whether this staining indicates specific or nonspecific binding of the anti-ChR1 polyclonal antiserum.

Growth in acetate affects ChR1 in min1 mutant cells. Chlamydomonas can grow photoautotrophically in the presence of light and CO2, mixotrophically in the presence of both acetate and light, and heterotrophically in the dark, utilizing acetate as a carbon source. In min1 nonsense mutant cells grown photoautotrophically (in acetate-free M medium), the eyespot pigment granule layers are relatively disorganized and are not apposed to the plasma membrane (33). Surprisingly, in min1 cells grown mixotrophically (in acetate-containing R medium), the pigment granule layers are more organized and are apposed to the plasma membrane (33). To determine whether increased organization of the granule layers is correlated with increased levels of ChR1, Western blots of total cellular protein from min1 nonsense mutant cells (strain 12-12) grown in the light, with or without acetate, were probed with anti-ChR1 (Fig. 8A). Contrary to what was expected in min1 cells grown with acetate (pigment granules more organized), the level of ChR1 was only 70% ± 10% of that in min1 cells grown without acetate (pigment granules disorganized). This difference was not observed in wild-type cells. Immunofluorescence analyses were consistent with the Western blot data (Fig. 8B); equatorial spots of ChR1 fluorescence were less frequent and noticeably smaller in min1 cells grown mixotrophically than in those grown photoautotrophically. Again, this difference was not observed in wild-type cells. These data suggest that ChR1 expression and/or stability is more sensitive to the absence of MIN1 in cells grown mixotrophically than in cells grown photoautotrophically.


Figure 8
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  		FIG. 8. ChR1 levels are very low in min1 cells grown in acetate-containing medium. (A) Western blot of total cellular protein from wild-type (wt) or min1 mutant (strain 12-12) cells grown in medium either lacking (M) or containing (R) acetate. The blot was probed with a polyclonal antibody against the photoreceptor ChR1 (5) followed by antitubulin (clone B-5-1-2; Sigma, St. Louis, MO). (B) Anti-ChR1 immunofluorescence of wild-type or min1 (strain 12-12) cells grown in either the absence (panels M) or presence (panels R) of acetate. For the min1 cells grown in M or R medium, both a field of cells and representative individual cells are shown. The arrows point to equatorial anti-ChR1; fluorescence at the anterior ends of the cells is most likely the result of nonspecific binding of anti-ChR1 in the region of the basal bodies (5).


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DISCUSSION
 
The eyespot of the green alga Chlamydomonas is a photosensory structure required for phototaxis. In min1 (mini-eyed) mutants, eyespot pigment granule numbers are reduced, and apposition of the plastid and plasma membrane components of the eyespot is disrupted (33). Here we describe the identification and initial characterization of the MIN1 gene and the encoded protein. The data highlight the novel domain composition of MIN1 and suggest that MIN1 promotes both apposition between the plastid and plasma membranes and expression and/or stability of the ChR1 photoreceptor.

The MIN1 protein contains N-terminal C2 (phospholipid membrane-binding) and C-terminal LysM (peptidoglycan-binding) domains, separated by a membrane-spanning {alpha}-helix. The LysM domain was originally found in bacterial cell wall-degrading enzymes that bind peptidoglycans (3). More recently the domain has been identified in plant proteins, specifically plasma membrane-localized receptors that interact with NOD (nodulation) factors in the cell walls of rhyzobial bacteria (37, 64, 76) and membrane proteins that trigger the defense response to chitin oligosaccharides in the cell walls of pathogenic fungi (27, 29). A LysM domain is also present in the Chlamydomonas eyespot protein, EYE2 (66, 71), and in a number of predicted Chlamydomonas proteins (unpublished observation), but potential interacting partners for these domains remain unknown. The MIN1 LysM domain is most similar to LysM domains in hypothetical proteins from the green alga Ostreococcus tauri and a variety of eubacteria. Each of these proteins also contains either a predicted membrane-spanning helix or a BON domain, which is hypothesized to bind to membranes (83). Together with proteomics data identifying MIN1 in eyespots (71) and the observation that MIN1 does not contain a chloroplast-targeting sequence identifiable by homology, the data suggest that MIN1 is embedded in the chloroplast envelope or plasma membrane in the eyespot.

The MIN1 C2 domain is also most similar to C2 domains in predicted membrane-associated proteins. C2 domains fold into an eight-stranded β-sheet "sandwich," and membrane association often requires Ca2+ coordination by aspartate residues in loops connecting individual β-strands. Membrane association of some C2 domains requires lysine residues on one side of the sandwich. A MIN1 C2 domain-YFP fusion protein localized to the chloroplast envelope in moss cells, and mutation of the two loop aspartate residues conserved in MIN1 abolished this association. These data indicate that the MIN1 C2 domain is capable of membrane association in a manner similar to that of more well-characterized C2 domains (34). However, Chlamydomonas transformants containing full-length MIN1 with the same aspartate residue mutations were phototactic and had wild-type eyespots. Similarly, transformants with mutations in conserved lysine residues in the MIN1 C2 domain were phototactic and had normal eyespots. One explanation for these data is that properties of the full-length MIN1 protein and/or the Chlamydomonas cellular environment minimize the requirement for these conserved residues. In previous studies (46, 75), the phenotypic consequences of C2 domain mutations were affected by the protein and cellular context, consistent with the hypothesis that interaction of C2 domains with phospholipid membranes is dependent on the distribution of electrostatic potential on the surface of the binding site rather than the presence of specific residues or Ca2+ coordination (49, 55). A second possibility is that the membrane-binding potential of the MIN1 C2 domain is not required for eyespot assembly. Either the C2 domain does not interact with membranes in Chlamydomonas, or the function of the domain in eyespot assembly is redundant. The existence of another protein that promotes eyespot membrane apposition could also explain the increased organization of the pigment granule/thylakoid membrane layers in min1 mutants grown mixotrophically. Further analyses are required to determine whether the MIN1 C2 domain associates with a Chlamydomonas membrane and, if so, whether the association is Ca2+ dependent and/or essential for eyespot assembly.

To date, the C2-plus-LysM domain composition is unique to MIN1, perhaps reflecting the fact that Chlamydomonas is the only eyespot-containing organism for which the complete genome sequence is available (48). The similarity of the C2 domain to eukaryotic sequences and of the LysM domain to eubacterial sequences prompts the speculation that MIN1 is the result of domain shuffling between genes encoding membrane-associated proteins in the original eukaryotic host and the endosymbiotic cyanobacterium and reflects the symbiotic origins of the eyespot (7, 19). Eyespot assembly requires that organization of the plastid pigment granule/thylakoid membrane layers, derived from an endosymbiotic cyanobacterium, is coordinated with localization of photoreceptors and other plasma membrane components, some of which are presumably derived from the host. How did this intricate coordination evolve? Insight may come from analyses of the function of a MIN1-like protein in modern symbiotic relationships such as the developing symbiosis between a green alga in the genus Nephroselmis and the flagellate Hatena arenicola (56, 57). Hatena cells containing an engulfed Nephroselmis cell have an apical eyespot in which Nephroselmis plastid and plasma membranes are apposed to the Hatena plasma membrane. At cell division, the Nephroselmis cell and the eyespot are inherited by one of the daughters, while the other daughter develops an apical feeding apparatus and resumes a phagocytic lifestyle. Are MIN1-like proteins encoded by the Hatena and/or Nephroselmis genome? If so, do they function in eyespot assembly in symbiotic cells and/or assembly of the feeding apparatus in phagocytic Hatena cells? Future molecular and cell biological studies should provide answers to these questions.

In min1 mutants grown photoautotrophically, the eyespot pigment granule/thylakoid membrane layers are disorganized and the overlying chloroplast envelope is no longer apposed to the plasma membrane (33). In cells grown mixotrophically in the light with acetate, min1 eyespots are more ordered and the chloroplast envelope and plasma membrane remain apposed. Do MIN1 and/or the physiological state of the cell also affect the plasma membrane components of the eyespot, specifically the photoreceptors? In min1 cells grown photoautotrophically, the level of ChR1, a rhodopsin family eyespot photoreceptor, is lower than that in wild-type cells, indicating that MIN1 promotes ChR1 expression and/or stability. ChR1 was even more reduced in min1 mutant cells, but not wild-type cells, grown mixotrophically, which suggests that under mixotrophic conditions, the requirement for MIN1 is more stringent despite the apparent increased organization of the plastid components of the eyespot.

In approximately 25% of min1 cells grown photoautotrophically, ChR1 was found in two or more roughly equatorial aggregations in the plasma membrane. This pattern is notably different from that of the pigment granules in photoautotrophically grown cells, which occur as a single aggregation that is disorganized and no longer apposed to the plasma membrane (33). This observation is consistent with the hypothesis that photoreceptor localization is not dependent solely on proper organization of the underlying pigment granule layers. We propose a testable model in which MIN1 is embedded in the plasma membrane or chloroplast envelope in the eyespot. The MIN1 C2 domain is predicted to promote membrane apposition, perhaps in combination with another eyespot protein. MIN1 also promotes expression and/or stabilization of ChR1; however, neither MIN1 nor proper organization of the pigment granule/thylakoid membrane layers is required for proper localization of the ChR1 photoreceptor.


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ACKNOWLEDGMENTS
 
We are grateful to Charles Quinton, Douglas Roberts, and Feliza Bourguet for the original identification of the min1 insertion strain, H6-2. We also sincerely thank Peter Hegemann for use of the ChR1 antibody, Samuel Ward for use of and assistance with the Leica microscope, Greg Pazour and George Witman for providing the Chlamydomonas cDNA library, and Patrice Hamel for providing strains 1B– and 4A+. Jacqueline Baca assisted with the Chlamydomonas transformations, Matt Cordes kindly provided instruction on the use of PyMol, and Derrick Sund and Joseph Boyd commented on the manuscript. Melissa Schonauer provided support throughout the course of this work.

This work was funded by NIH grant GM60933 to C.L.D.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biochemistry and Molecular Biophysics, University of Arizona, Life Sciences South no. 454, 1007 E. Lowell Street, Tucson, AZ 85721-0106. Phone: (520) 621-3569. Fax: (520) 621-3709. E-mail: telsa{at}email.arizona.edu Back

{triangledown} Published ahead of print on 10 October 2008. Back

{dagger} Supplemental material for this article may be found at http://ec.asm.org/. Back


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REFERENCES
 
    1
  1. Altschul, S. F., W. Gish, W. Miller, E. W. Myers, and D. J. Lipman. 1990. Basic local alignment search tool. J. Mol. Biol. 215:403-410.[CrossRef][Medline]
    2. 2
  2. Auchincloss, A. H., A. I. Loroch, and J. D. Rochaix. 1999. The argininosuccinate lyase gene of Chlamydomonas reinhardtii: cloning of the cDNA and its characterization as a selectable shuttle marker. Mol. Gen. Genet. 261:21-30.[CrossRef][Medline]
    3. 3
  3. Bateman, A., and M. Bycroft. 2000. The structure of a LysM domain from E. coli membrane-bound lytic murein transglycosylase D (MltD). J. Mol. Biol. 299:1113-1119.[CrossRef][Medline]
    4. 4
  4. Bateman, A., L. Coin, R. Durbin, R. D. Finn, V. Hollich, S. Griffiths-Jones, A. Khanna, M. Marshall, S. Moxon, E. L. Sonnhammer, D. J. Studholme, C. Yeats, and S. R. Eddy. 2004. The Pfam protein families database. Nucleic Acids Res. 32:D138-D141.[Abstract/Free Full Text]
    5. 5
  5. Berthold, P., S. P. Tsunoda, O. P. Ernst, W. Mages, D. Gradmann, and P. Hegemann. 13 June 2008. Channelrhodopsin-1 initiates phototaxis and photophobic responses in Chlamydomonas by immediate light-induced depolarization. Plant Cell 20:1655-1677. [Epub ahead of print.]
    6. 6
  6. Bielnicki, J., Y. Devedjiev, U. Derewenda, Z. Dauter, A. Joachimiak, and Z. S. Derewenda. 2006. B. subtilis ykuD protein at 2.0 A resolution: insights into the structure and function of a novel, ubiquitous family of bacterial enzymes. Proteins 62:144-151.[CrossRef][Medline]
    7. 7
  7. Cavalier-Smith, T. 2000. Membrane heredity and early chloroplast evolution. Trends Plant Sci. 5:174-182.[CrossRef][Medline]
    8. 8
  8. Cerutti, H., A. M. Johnson, N. W. Gillham, and J. E. Boynton. 1997. Epigenetic silencing of a foreign gene in nuclear transformants of Chlamydomonas. Plant Cell 9:925-945.[Abstract/Free Full Text]
    9. 9
  9. Cho, W. 2001. Membrane targeting by C1 and C2 domains. J. Biol. Chem. 276:32407-32410.[Free Full Text]
    10. 10
  10. DeLano, W. L. 2002. The PyMOL molecular graphics system. DeLano Scientific, Palo Alto, CA.
    11. 11
  11. Dieckmann, C. L. 2003. Eyespot placement and assembly in the green alga Chlamydomonas. Bioessays 25:410-416.[CrossRef][Medline]
    12. 12
  12. Doerks, T., M. Strauss, M. Brendel, and P. Bork. 2000. GRAM, a novel domain in glucosyltransferases, myotubularins and other putative membrane-associated proteins. Trends Biochem. Sci. 25:483-485.[CrossRef][Medline]
    13. 13
  13. Dunahay, T. G. 1993. Transformation of Chlamydomonas reinhardtii with silicon carbide whiskers. BioTechniques 15:452-460.
    14. 14
  14. Dutcher, S. K. 2000. Chlamydomonas reinhardtii: biological rationale for genomics. J. Eukaryot. Microbiol. 47:340-349.[CrossRef][Medline]
    15. 15
  15. Dyall, S. D., M. T. Brown, and P. J. Johnson. 2004. Ancient invasions: from endosymbionts to organelles. Science 304:253-257.[Abstract/Free Full Text]
    16. 16
  16. Fernandez, I., D. Araç, J. Ubach, S. H. Gerber, O. Shin, Y. Gao, R. G. Anderson, T. C. Südhof, and J. Rizo. 2001. Three-dimensional structure of the synaptotagmin 1 C2B-domain: synaptotagmin 1 as a phospholipid binding machine. Neuron 32:1057-1069.[CrossRef][Medline]
    17. 17
  17. Field, J., J. Nikawa, D. Broek, B. MacDonald, L. Rodgers, I. A. Wilson, R. A. Lerner, and M. Wigler. 1988. Purification of a RAS-responsive adenylyl cyclase complex from Saccharomyces cerevisiae by use of an epitope addition method. Mol. Cell. Biol. 8:2159-2165.[Abstract/Free Full Text]
    18. 18
  18. Foster, K. W., and R. D. Smyth. 1980. Light antennas in phototactic algae. Microbiol. Rev. 44:572-630.[Free Full Text]
    19. 19
  19. Gehring, W. J. 2005. New perspectives on eye development and the evolution of eyes and photoreceptors. J. Hered. 96:171-184.[Abstract/Free Full Text]
    20. 20
  20. Goldschmidt-Clermont, M., and M. Rahire. 1986. Sequence, evolution and differential expression of the two genes encoding variant small subunits of ribulose bisphosphate carboxylase/oxygenase in Chlamydomonas reinhardtii. J. Mol. Biol. 191:421-432.[CrossRef][Medline]
    21. 21
  21. Gorman, D. S., and R. P. Levine. 1965. Cytochrome f and plastocyanin: their sequence in the photosynthetic electron transport chain of Chlamydomonas reinhardi. Proc. Natl. Acad. Sci. USA 54:1665-1669.[Free Full Text]
    22. 22
  22. Harris, E. H. 1989. The Chlamydomonas sourcebook. Academic Press, San Diego, CA.
    23. 23
  23. Harris, E. H. 2001. Chlamydomonas as a model organism. Annu. Rev. Plant Physiol. Plant Mol. Biol. 52:363-406.[CrossRef][Medline]
    24. 24
  24. Hegemann, P., K. Neumeier, U. Hegemann, and E. Kuehnle. 1990. The role of calcium in Chlamydomonas photomovement responses as analysed by calcium channel inhibitors. Photochem. Photobiol. 52:575-583.[Medline]
    25. 25
  25. Holmes, J. A., and S. K. Dutcher. 1989. Cellular asymmetry in Chlamydomonas reinhardtii. J. Cell Sci. 94:273-285.[Abstract/Free Full Text]
    26. 26
  26. Jiménez, J. L., G. R. Smith, B. Contreras-Moreira, J. G. Sgouros, F. A. Meunier, P. A. Bates, and G. Schiavo. 2003. Functional recycling of C2 domains throughout evolution: a comparative study of synaptotagmin, protein kinase C and phospholipase C by sequence, structural and modelling approaches. J. Mol. Biol. 333:621-639.[CrossRef][Medline]
    27. 27
  27. Kaku, H., Y. Nishizawa, N. Ishii-Minami, C. Akimoto-Tomiyama, N. Dohmae, K. Takio, E. Minami, and N. Shibuya. 2006. Plant cells recognize chitin fragments for defense signaling through a plasma membrane receptor. Proc. Natl. Acad. Sci. USA 103:11086-11091.[Abstract/Free Full Text]
    28. 28
  28. Kelley, L. A., R. M. MacCallum, and M. J. Sternberg. 2000. Enhanced genome annotation using structural profiles in the program 3D-PSSM. J. Mol. Biol. 299:499-520.[Medline]
    29. 29
  29. Knogge, W., and D. Scheel. 2006. LysM receptors recognize friend and foe. Proc. Natl. Acad. Sci. USA 103:10829-10830.[Free Full Text]
    30. 30
  30. Kreimer, G., and M. Melkonian. 1990. Reflection confocal laser scanning microscopy of eyespots in flagellated green algae. Eur. J. Cell Biol. 53:101-111.[Medline]
    31. 31
  31. Krogh, A., B. Larsson, G. von Heijne, and E. L. Sonnhammer. 2001. Predicting transmembrane protein topology with a hidden Markov model: application to complete genomes. J. Mol. Biol. 305:567-580.[CrossRef][Medline]
    32. 32
  32. Laemmli, U. K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685.[CrossRef][Medline]
    33. 33
  33. Lamb, M. R., S. K. Dutcher, C. K. Worley, and C. L. Dieckmann. 1999. Eyespot-assembly mutants in Chlamydomonas reinhardtii. Genetics 153:721-729.[Abstract/Free Full Text]
    34. 34
  34. Lemmon, M. A. 2008. Membrane recognition by phospholipid-binding domains. Nat. Rev. Mol. Cell Biol. 9:99-111.[CrossRef][Medline]
    35. 35
  35. Letunic, I., L. Goodstadt, N. J. Dickens, T. Doerks, J. Schultz, R. Mott, F. Ciccarelli, R. R. Copley, C. P. Ponting, and P. Bork. 2002. Recent improvements to the SMART domain-based sequence annotation resource. Nucleic Acids Res. 30:242-244.[Abstract/Free Full Text]
    36. 36
  36. Levitan, A., T. Trebitsh, V. Kiss, Y. Pereg, I. Dangoor, and A. Danon. 2005. Dual targeting of the protein disulfide isomerase RB60 to the chloroplast and the endoplasmic reticulum. Proc. Natl. Acad. Sci. USA 102:6225-6230.[Abstract/Free Full Text]
    37. 37
  37. Limpens, E., C. Franken, P. Smit, J. Willemse, T. Bisseling, and R. Geurts. 2003. LysM domain receptor kinases regulating rhizobial Nod factor-induced infection. Science 302:630-633.[Abstract/Free Full Text]
    38. 38
  38. Liu, L., X. Song, D. He, C. Komma, A. Kita, J. V. Virbasius, G. Huang, H. D. Bellamy, K. Miki, M. P. Czech, and G. W. Zhou. 2006. Crystal structure of the C2 domain of class II phosphatidylinositide 3-kinase C2alpha. J. Biol. Chem. 281:4254-4260.[Abstract/Free Full Text]
    39. 39
  39. Lukashin, A. V., and M. Borodovsky. 1998. GeneMark.hmm: new solutions for gene finding. Nucleic Acids Res. 26:1107-1115.[Abstract/Free Full Text]
    40. 40
  40. Lumbreras, V., D. R. Stevens, and S. Purton. 1998. Efficient foreign gene expression in Chlamydomonas reinhardtii mediated by an endogenous intron. Plant J. 14:441-447.[CrossRef]
    41. 41
  41. Madden, T. L., R. L. Tatusov, and J. Zhang. 1996. Applications of network BLAST server. Methods Enzymol. 266:131-141.[Medline]
    42. 42
  42. Malmberg, N. J., S. Varma, E. Jakobsson, and J. J. Falke. 2004. Ca2+ activation of the cPLA2 C2 domain: ordered binding of two Ca2+ ions with positive cooperativity. Biochemistry 43:16320-16328.
    43. 43
  43. Marchler-Bauer, A., J. B. Anderson, C. DeWeese-Scott, N. D. Fedorova, L. Y. Geer, S. He, D. I. Hurwitz, J. D. Jackson, A. R. Jacobs, C. J. Lanczycki, C. A. Liebert, C. Liu, T. Madej, G. H. Marchler, R. Mazumder, A. N. Nikolskaya, A. R. Panchenko, B. S. Rao, B. A. Shoemaker, V. Simonyan, J. S. Song, P. A. Thiessen, S. Vasudevan, Y. Wang, R. A. Yamashita, J. J. Yin, and S. H. Bryant. 2003. CDD: a curated Entrez database of conserved domain alignments. Nucleic Acids Res. 31:383-387.[Abstract/Free Full Text]
    44. 44
  44. Melkonian, M., and H. Robenek. 1980. Eyespot membranes of Chlamydomonas reinhardii: a freeze-fracture study. J. Ultrastruct. Res. 72:90-102.[CrossRef][Medline]
    45. 45
  45. Meller, J., and R. Elber. 2001. Linear programming optimization and a double statistical filter for protein threading protocols. Proteins 45:241-261.[CrossRef][Medline]
    46. 46
  46. Melowic, H. R., R. V. Stahelin, N. R. Blatner, W. Tian, K. Hayashi, A. Altman, and W. Cho. 2007. Mechanism of diacylglycerol-induced membrane targeting and activation of protein kinase Ctheta. J. Biol. Chem. 282:21467-21476.[Abstract/Free Full Text]
    47. 47
  47. Merchant, S., and L. Bogorad. 1987. The Cu(II)-repressible plastidic cytochrome c. Cloning and sequence of a complementary DNA for the pre-apoprotein. J. Biol. Chem. 262:9062-9067.[Abstract/Free Full Text]
    48. 48
  48. Merchant, S. S., S. E. Prochnik, O. Vallon, E. H. Harris, S. J. Karpowicz, G. B. Witman, A. Terry, A. Salamov, L. K. Fritz-Laylin, L. Marechal-Drouard, W. F. Marshall, L. H. Qu, D. R. Nelson, A. A. Sanderfoot, M. H. Spalding, V. V. Kapitonov, Q. Ren, P. Ferris, E. Lindquist, H. Shapiro, S. M. Lucas, J. Grimwood, J. Schmutz, P. Cardol, H. Cerutti, G. Chanfreau, C. L. Chen, V. Cognat, M. T. Croft, R. Dent, S. Dutcher, E. Fernandez, H. Fukuzawa, D. Gonzalez-Ballester, D. Gonzalez-Halphen, A. Hallmann, M. Hanikenne, M. Hippler, W. Inwood, K. Jabbari, M. Kalanon, R. Kuras, P. A. Lefebvre, S. D. Lemaire, A. V. Lobanov, M. Lohr, A. Manuell, I. Meier, L. Mets, M. Mittag, T. Mittelmeier, J. V. Moroney, J. Moseley, C. Napoli, A. M. Nedelcu, K. Niyogi, S. V. Novoselov, I. T. Paulsen, G. Pazour, S. Purton, J. P. Ral, D. M. Riano-Pachon, W. Riekhof, L. Rymarquis, M. Schroda, D. Stern, J. Umen, R. Willows, N. Wilson, S. L. Zimmer, J. Allmer, J. Balk, K. Bisova, C. J. Chen, M. Elias, K. Gendler, C. Hauser, M. R. Lamb, H. Ledford, J. C. Long, J. Minagawa, M. D. Page, J. Pan, W. Pootakham, S. Roje, A. Rose, E. Stahlberg, A. M. Terauchi, P. Yang, S. Ball, C. Bowler, C. L. Dieckmann, V. N. Gladyshev, P. Green, R. Jorgensen, S. Mayfield, B. Mueller-Roeber, S. Rajamani, R. T. Sayre, P. Brokstein, I. Dubchak, D. Goodstein, L. Hornick, Y. W. Huang, J. Jhaveri, Y. Luo, D. Martinez, W. C. Ngau, B. Otillar, A. Poliakov, A. Porter, L. Szajkowski, G. Werner, K. Zhou, I. V. Grigoriev, D. S. Rokhsar, and A. R. Grossman. 2007. The Chlamydomonas genome reveals the evolution of key animal and plant functions. Science 318:245-250.[Abstract/Free Full Text]
    49. 49
  49. Murray, D., and B. Honig. 2002. Electrostatic control of the membrane targeting of C2 domains. Mol. Cell 9:145-154.[CrossRef][Medline]
    50. 50
  50. Nagashima, T., F. Hayashi, and S. Yokoyama. 2004. C2 domain-containing protein from putative elicitor-responsive gene. RIKEN Structural Genomics/Proteomics Initiative. RIKEN, Yokohama, Japan.
    51. 51
  51. Nagel, G., D. Ollig, M. Fuhrmann, S. Kateriya, A. M. Musti, E. Bamberg, and P. Hegemann. 2002. Channelrhodopsin-1: a light-gated proton channel in green algae. Science 296:2395-2398.[Abstract/Free Full Text]
    52. 52
  52. Nagel, G., T. Szellas, W. Huhn, S. Kateriya, N. Adeishvili, P. Berthold, D. Ollig, P. Hegemann, and E. Bamberg. 2003. Channelrhodopsin-2, a directly light-gated cation-selective membrane channel. Proc. Natl. Acad. Sci. USA 100:13940-13945.[Abstract/Free Full Text]
    53. 53
  53. Nalefski, E. A., and J. J. Falke. 1996. The C2 domain calcium-binding motif: structural and functional diversity. Protein Sci. 5:2375-2390.[Medline]
    54. 54
  54. Naylor, C. E., J. T. Eaton, A. Howells, N. Justin, D. S. Moss, R. W. Titball, and A. K. Basak. 1998. Structure of the key toxin in gas gangrene. Nat. Struct. Biol. 5:738-746.[CrossRef][Medline]
    55. 55
  55. Ochoa, W. F., J. Garcia-Garcia, I. Fita, S. Corbalán-Garcia, N. Verdaguer, and J. C. Gómez-Fernández. 2001. Structure of the C2 domain from novel protein kinase Cepsilon. A membrane binding model for Ca(2+)-independent C2 domains. J. Mol. Biol. 311:837-849.[CrossRef][Medline]
    56. 56
  56. Okamoto, N., and I. Inouye. 2005. A secondary symbiosis in progress? Science 310:287.[Abstract/Free Full Text]
    57. 57
  57. Okamoto, N., and I. Inouye. 2006. Hatena arenicola gen. et sp. nov., a katablepharid undergoing probable plastid acquisition. Protist 157:401-419.[Medline]
    58. 58
  58. Pazour, G. J., O. A. Sineshchekov, and G. B. Witman. 1995. Mutational analysis of the phototransduction pathway of Chlamydomonas reinhardtii. J. Cell Biol. 131:427-440.[Abstract/Free Full Text]
    59. 59
  59. Peers, G., and K. K. Niyogi. 2008. Pond scum genomics: the genomes of Chlamydomonas and Ostreococcus. Plant Cell 20:502-507.[Free Full Text]
    60. 60
  60. Perisic, O., S. Fong, D. E. Lynch, M. Bycroft, and R. L. Williams. 1998. Crystal structure of a calcium-phospholipid binding domain from cytosolic phospholipase A2. J. Biol. Chem. 273:1596-1604.[Abstract/Free Full Text]
    61. 61
  61. Pidcock, E., and G. R. Moore. 2001. Structural characteristics of protein binding sites for calcium and lanthanide ions. J. Biol. Inorg. Chem. 6:479-489.[CrossRef][Medline]
    62. 62
  62. Ponting, C. P., and P. J. Parker. 1996. Extending the C2 domain family: C2s in PKCs delta, epsilon, eta, theta, phospholipases, GAPs, and perforin. Protein Sci. 5:162-166.[Medline]
    63. 63
  63. Purton, S., and J.-D. Rochaix. 1994. Complementation of a Chlamydomonas reinhardtii mutant using a genomic cosmid library. Plant Mol. Biol. 24:533-537.[CrossRef][Medline]
    64. 64
  64. Radutoiu, S., L. H. Madsen, E. B. Madsen, A. Jurkiewicz, E. Fukai, E. M. Quistgaard, A. S. Albrektsen, E. K. James, S. Thirup, and J. Stougaard. 2007. LysM domains mediate lipochitin-oligosaccharide recognition and Nfr genes extend the symbiotic host range. EMBO J. 26:3923-3935.[CrossRef][Medline]
    65. 65
  65. Rizo, J., and T. C. Südhof. 1998. C2-domains, structure and function of a universal Ca2+-binding domain. J. Biol. Chem. 273:15879-15882.[Free Full Text]
    66. 66
  66. Roberts, D. G., M. R. Lamb, and C. L. Dieckmann. 2001. Characterization of the EYE2 gene required for eyespot assembly in Chlamydomonas reinhardtii. Genetics 158:1037-1049.[Abstract/Free Full Text]
    67. 67
  67. Rodríguez-Alfaro, J. A., J. C. Gómez-Fernández, and S. Corbalán-Garcia. 2004. Role of the lysine-rich cluster of the C2 domain in the phosphatidylserine-dependent activation of PKCalpha. J. Mol. Biol. 335:1117-1129.[CrossRef][Medline]
    68. 68
  68. Rohr, J., N. Sarkar, S. Balenger, B. R. Jeong, and H. Cerutti. 2004. Tandem inverted repeat system for selection of effective transgenic RNAi strains in Chlamydomonas. Plant J. 40:611-621.[CrossRef][Medline]
    69. 69
  69. Ruffer, U., and W. Nultsch. 1998. Flagellar coordination in Chlamydomonas cells held on micropipettes. Cell Motil. Cytoskel. 41:297-307.[Medline]
    70. 70
  70. Sánchez-Bautista, S., C. Marín-Vicente, J. C. Gómez-Fernández, and S. Corbalán-Garcia. 2006. The C2 domain of PKCalpha is a Ca2+-dependent PtdIns(4,5)P2 sensing domain: a new insight into an old pathway. J. Mol. Biol. 362:901-914.[CrossRef][Medline]
    71. 71
  71. Schmidt, M., G. Gessner, M. Luff, I. Heiland, V. Wagner, M. Kaminski, S. Geimer, N. Eitzinger, T. Reissenweber, O. Voytsekh, M. Fiedler, M. Mittag, and G. Kreimer. 2006. Proteomic analysis of the eyespot of Chlamydomonas reinhardtii provides novel insights into its components and tactic movements. Plant Cell 18:1908-1930.[Abstract/Free Full Text]
    72. 72
  72. Schultz, J., F. Milpetz, P. Bork, and C. P. Ponting. 1998. SMART, a simple modular architecture research tool: identification of signaling domains. Proc. Natl. Acad. Sci. USA 95:5857-5864.[Abstract/Free Full Text]
    73. 73
  73. Silflow, C. D., R. L. Chisholm, T. W. Conner, and L. P. Ranum. 1985. The two alpha-tubulin genes of Chlamydomonas reinhardtii code for slightly different proteins. Mol. Cell. Biol. 5:2389-2398.[Abstract/Free Full Text]
    74. 74
  74. Sineshchekov, O. A., K. H. Jung, and J. L. Spudich. 2002. Two rhodopsins mediate phototaxis to low- and high-intensity light in Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. USA 99:8689-8694.[Abstract/Free Full Text]
    75. 75
  75. Six, D. A., and E. A. Dennis. 2003. Essential Ca(2+)-independent role of the group IVA cytosolic phospholipase A(2) C2 domain for interfacial activity. J. Biol. Chem. 278:23842-23850.[Abstract/Free Full Text]
    76. 76
  76. Spaink, H. P. 2004. Specific recognition of bacteria by plant LysM domain receptor kinases. Trends Microbiol. 12:201-204.[CrossRef][Medline]
    77. 77
  77. Steen, A., G. Buist, K. J. Leenhouts, M. El Khattabi, F. Grijpstra, A. L. Zomer, G. Venema, O. P. Kuipers, and J. Kok. 2003. Cell wall attachment of a widely distributed peptidoglycan binding domain is hindered by cell wall constituents. J. Biol. Chem. 278:23874-23881.[Abstract/Free Full Text]
    78. 78
  78. Suzuki, T., K. Yamasaki, S. Fujita, K. Oda, M. Iseki, K. Yoshida, M. Watanabe, H. Daiyasu, H. Toh, E. Asamizu, S. Tabata, K. Miura, H. Fukuzawa, S. Nakamura, and T. Takahashi. 2003. Archaeal-type rhodopsins in Chlamydomonas: model structure and intracellular localization. Biochem. Biophys. Res. Commun. 301:711-717.[CrossRef][Medline]
    79. 79
  79. Tam, L. W., and P. A. Lefebvre. 1993. Cloning of flagellar genes in Chlamydomonas reinhardtii by DNA insertional mutagenesis. Genetics 135:375-384.[Abstract]
    80. 80
  80. Thompson, J. D., D. G. Higgins, and T. J. Gibson. 1994. Improved sensitivity of profile searches through the use of sequence weights and gap excision. Comput. Appl. Biosci. 10:19-29.[Abstract/Free Full Text]
    81. 81
  81. Tobi, D., and R. Elber. 2000. Distance-dependent, pair potential for protein folding: results from linear optimization. Proteins 41:40-46.[Medline]
    82. 82
  82. Witman, G. B. 1993. Chlamydomonas phototaxis. Trends Cell Biol. 3:403-408.[CrossRef][Medline]
    83. 83
  83. Yeats, C., and A. Bateman. 2003. The BON domain: a putative membrane-binding domain. Trends Biochem. Sci. 28:352-355.[CrossRef][Medline]

Eukaryotic Cell, December 2008, p. 2100-2112, Vol. 7, No. 12
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