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Journal of Bacteriology, November 2004, p. 7420-7428, Vol. 186, No. 21
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.21.7420-7428.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

AplA, a Member of a New Class of Phycobiliproteins Lacking a Traditional Role in Photosynthetic Light Harvesting

Beronda L. Montgomery,1 Elena Silva Casey,2 Arthur R. Grossman,3 and David M. Kehoe1*

Department of Biology, Indiana University, Bloomington, Indiana,1 Department of Biology, Georgetown University, Washington, D.C.,2 Department of Plant Biology, Carnegie Institution of Washington, Stanford, California3

Received 24 March 2004/ Accepted 2 August 2004


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ABSTRACT
 
All known phycobiliproteins have light-harvesting roles during photosynthesis and are found in water-soluble phycobilisomes, the light-harvesting complexes of cyanobacteria, cyanelles, and red algae. Phycobiliproteins are chromophore-bearing proteins that exist as heterodimers of {alpha} and ß subunits, possess a number of highly conserved amino acid residues important for dimerization and chromophore binding, and are invariably 160 to 180 amino acids long. A new and unusual group of proteins that is most closely related to the allophycocyanin members of the phycobiliprotein superfamily has been identified. Each of these proteins, which have been named allophycocyanin-like (Apl) proteins, apparently contains a 28-amino-acid extension at its amino terminus relative to allophycocyanins. Apl family members possess the residues critical for chromophore interactions, but substitutions are present at positions implicated in maintaining the proper {alpha}-ß subunit interactions and tertiary structure of phycobiliproteins, suggesting that Apl proteins are able to bind chromophores but fail to adopt typical allophycocyanin conformations. AplA isolated from the cyanobacterium Fremyella diplosiphon contained a covalently attached chromophore and, although present in the cell under a number of conditions, was not detected in phycobilisomes. Thus, Apl proteins are a new class of photoreceptors with a different cellular location and structure than any previously described members of the phycobiliprotein superfamily.


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INTRODUCTION
 
During photosynthesis, the efficiency of light energy capture is increased by the presence of light-harvesting structures, and a wide variety exists in oxygen-evolving photosynthetic organisms (25). Arguably the best understood of these structures are phycobilisomes (PBS), the light-harvesting complexes of cyanobacteria, cyanelles, and red algae (19, 23, 44). PBS collect and transfer light energy to integral membrane proteins of photosynthetic reaction centers and consist of two distinct regions, a core and a region of peripheral rods. Most PBS contain four to eight peripheral rods, which are cylindrical structures that project from the core. Peripheral rods are responsible for light energy capture and unidirectional energy transfer to the core. A PBS core generally contains two or three cylinders that are perpendicular to their linked peripheral rods. One face of the core associates with the thylakoid membrane, allowing energy transfer from peripheral rods to the photosynthetic reaction centers.

Both the peripheral rods and the core contain two classes of proteins. Phycobiliproteins have one or more covalently attached open-chain tetrapyrrole (bilin) chromophores and harvest most of the light energy. Linker peptides are paired with specific phycobiliproteins, have structural and energy transfer roles in PBS, and are predominantly not chromophorylated (49). Many different types of phycobiliproteins have been identified (3). Each type maximally absorbs light at a unique wavelength and is positioned in a specific location within the PBS. In the filamentous cyanobacterium Fremyella diplosiphon, the rods may contain phycoerythrin (PE; maximum absorption [{lambda}max], 560 nm) and three different types of phycocyanin (PC; {lambda}max, 620 nm), while the core is made up primarily of allophycocyanin (AP; {lambda}max, 650 nm) (48). All major species of phycobiliproteins are heterodimers consisting of {alpha} and ß subunits and are encoded by genes which usually are in an operon. Phycobiliprotein {alpha} and ß subunits are also uniform in length, with 160 to 180 amino acid (aa) residues, and X-ray crystallographic studies have shown that they all adopt similar three-dimensional structures (6).

Detailed analyses of PBS composition have uncovered three unorthodox phycobiliproteins in cyanobacteria. All are variants of the major AP {alpha} and ß subunit species found in PBS cores, which are encoded by the apcAB operon. The first to be discovered, AP B ({alpha}APB) (24), was most closely related in sequence to AP {alpha} subunits and determined to be a minor component of PBS cores (37, 38, 46) (NCBI accession no. AF059337). The {alpha}APB protein of Calothrix sp. strain PCC 7601, encoded by apcD, had an apparent length of 161 aa and was about 3% of the total core phycobiliprotein (29). apcD was unusual because no contiguous gene encoding an AP ß subunit was identified. Studies of Synechococcus sp. strain PCC 6301 revealed that APB dimerized with the same AP ß subunit that dimerized with the major AP {alpha} subunit (36). Although the role of {alpha}APB is not yet completely defined and PBS are assembled in its absence, {alpha}APB appears to be involved in transfer of energy to the photosynthetic reaction centers and partitioning of energy between photosystems I and II (20, 39, 52). The second unusual phycobiliprotein to be discovered was an 18.3-kDa peptide named ß18, also identified as a minor constituent within the core of the PBS of Synechococcus sp. strain PCC 6301 (37, 38, 51). ß18, which is similar to the AP ß subunit, is encoded by apcF, which has no AP subunit-encoding genes nearby in Synechococcus sp. strain PCC 7002 (20). Inactivation of apcF in this species did not lead to the loss of PBS assembly and had minor effects on PBS function. The doubling times of this mutant increased significantly during growth in green light (GL), and low-temperature fluorescence emission spectral data suggested impairment in energy transfer to photosystem II (52). A gene encoding a third unusual phycobiliprotein with similarity to AP {alpha} subunits was cloned from Calothrix sp. strain PCC 7601 and named apcA2. It was also not located near a gene encoding a ß subunit, and no information is available regarding its expression or the location or function of ApcA2, which has a predicted length of 160 aa (28).

Here we describe aplA (named for allophycocyanin-like A) from F. diplosiphon, which encodes a protein with sequence relatedness to AP family members. Phylogenetic analysis demonstrates that AplA, along with previously undescribed proteins from the cyanobacteria Nostoc punctiforme and Trichodesmium erythraeum, form a new and distinct class of unusual AP-like proteins that are not cotranscribed with complementary AP subunit partners and not closely related to the unorthodox phycobiliproteins described above. In addition, these proteins lack amino acids identified as important for normal phycobiliprotein structure and interaction with partner AP subunits and linker peptides but possess many residues critical for chromophore attachment and coordination. AplA is expressed in cells and, like other phycobiliproteins, has a covalently attached bilin chromophore. However, unlike any previously described phycobiliprotein, AplA is not found within PBS and thus appears to have a unique photoreception function.


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MATERIALS AND METHODS
 
Cyanobacterial growth conditions. F. diplosiphon (Calothrix or Tolypothrix sp. strain PCC 7601) cultures were grown as previously described (42). The wild type was Fd33 (also named SF33) (13), an F. diplosiphon mutant with normal photoresponses that forms discrete colonies on plates. Cells used for protein studies were harvested at mid-exponential phase of growth.

Bioinformatics. Sequence similarity analyses were performed using the BLAST program (2) of the National Center for Biotechnology Information (NCBI). Conventional single-letter amino acid designations were used. AplA isoelectric point and mass calculations were made using the PeptideMass tool (http://us.expasy.org/tools/peptide-mass.html). Protein family database analysis (5) was conducted by use of the Pfam database (http://pfam.wustl.edu/). Helix-turn-helix predictions were made by helix-turn-helix analysis (15), and multiple sequence alignments were performed with ClustalW (50) at the San Diego Supercomputing Center's Biology WorkBench site (http://workbench.sdsc.edu/). The ProtML (protein maximum likelihood) program was used to generate an unrooted tree with bootstrap values for phylogenetic comparisons (1).

Protein fold recognition was performed using Fischer's Hybrid Fold Recognition server (http://www.cs.bgu.ac.il/~bioinbgu/) (17), and sequence-structure homology recognition was done by FUGUE analysis (http://www-cryst.bioc.cam.ac.uk/fugue/) (43). The AplA three-dimensional structure was modeled based on structural similarity to the AP {alpha} chain of Spirulina platensis (NCBI Protein Database [PDB] entry 1alla) (9) by comparative protein modeling using a FUGUE output file and the RasMac PDB viewer (version 2.5).

DNA manipulation and antibody production. Standard approaches were used for cloning, sequencing, electrophoresis, and PCR amplifications (41). Two clones containing part or all of aplA were independently isolated from an F. diplosiphon genomic DNA library (33). The first, pDK1, complemented an F. diplosiphon black pigmentation (FdBk) mutant (32), and the second, pEC99, suppressed blue pigmentation mutants in F. diplosiphon (11). In both of these studies, aplA was designated ORFIII. To overproduce AplA for anti-AplA antibody production, the genomic DNA region containing aplA was cloned into pQE60 (QIAGEN, Valencia, Calif.) as follows. The primers APSP1 (5' CGGGATCCGAAGTGACTCAGGCACAATTA 3') and APSP2 (5' CGGGATCCAACCATTGCTTGCTGCTCTG 3'), each with a 5'-end BamHI site (underlined), were used to PCR amplify aplA. The PCR product was gel purified and cloned into the BamHI site of pQE60 to make pQE60APL, and the insert DNA and ligation junctions were sequenced. This added the sequence MGGS to the N terminus of AplA and a six-histidine extension to the carboxyl terminus. Escherichia coli M15 cells carrying pQE60APL were grown to early exponential phase in Luria-Bertani medium (41) with 100 µg of ampicillin/ml and 25 µg of kanamycin/ml and then were induced to synthesize high levels of AplA-His by addition of IPTG (isopropyl ß-D-thiogalactopyranoside; 2 mM final concentration) and subsequently grown at 37°C for 2 h prior to harvest. Cells were centrifuged at 4,000 x g for 10 min and then resuspended in 15 ml of ice-cold buffer A (50 mM Tris-HCl [pH 7.7], 300 mM NaCl, 10 µg of RNase A/ml), passed twice through a small French pressure cell at 4°C at 18,000 lb/in2, and then centrifuged at 12,000 x g for 20 min. A 3.5-ml volume of Talon resin (Clontech, Palo Alto, Calif.) was added to the decanted supernatant, which was then incubated overnight at 4°C with gentle shaking. The solution was loaded onto a Poly-Prep column (Bio-Rad Laboratories, Hercules, Calif.) at 4°C, and the resin was washed with 60 ml of buffer A and then with 30 ml of buffer B (50 mM phosphate buffer [pH 7.7], 300 mM NaCl, 10% glycerol). AplA-His was eluted using 15 ml of buffer B containing 0.5 M imidazole. Fractions containing AplA-His were pooled, and the imidazole was removed by a series of washes with buffer B in a Centricon 10 column (Millipore Corp., Bedford, Mass.). AplA-His was isolated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and used as the antigen for antibody production in rabbits (Cocalico Biologicals, Reamstown, Pa.). Anti-AplA antibody was affinity purified against AplA-His separated by SDS-PAGE.

Protein extraction, immunoblot, and zinc blot analyses. Mid-exponential-phase cultures of F. diplosiphon cells were chilled to 5°C in liquid nitrogen and centrifuged at 5,000 x g for 10 min at 4°C. Cell pellets were washed in the original culture volume of detergent-free extraction buffer (50 mM Tris-HCl [pH 7.5], 10% glycerol, 300 mM NaCl, 5 mM imidazole, 1 mM phenylmethylsulfonyl fluoride [PMSF], 1 mM 2-mercaptoethanol, 0.2 mM benzamidine, 50 µM 6-amino-n-hexanoic acid), and the pellet was collected as described above. Washed cells were resuspended in 1/20 of the original culture volume of extraction buffer and passed twice through a small French pressure cell at 18,000 lb/in2 at 4°C. The soluble fraction was separated by centrifuging it for 1 h at 100,000 x g at 4°C. The pellet containing the insoluble protein fraction was resuspended in extraction buffer containing 0.1% SDS. PBS were purified on sucrose gradients as previously described (14). Protein concentrations were determined using Bradford reagent (Bio-Rad Laboratories) with bovine serum albumin as the standard.

F. diplosiphon cells (100 ml) were grown to mid-exponential phase and harvested as described above, and AplA was immunoprecipitated using anti-AplA antibodies at 4°C as follows. Cell pellets were washed once with lysis buffer (50 mM Tris-HCl [pH 7.5], 150 mM NaCl, 0.5% [wt/vol] Triton X-100, 1 mM PMSF, 0.2 mM benzamidine, 50 µM 6-amino-n-hexanoic acid), recentrifuged, resuspended in 1.5 ml of lysis buffer, and passed twice through a French pressure cell at 18,000 lb/in2. Broken cells were centrifuged for 20 min at 16,000 x g, 1 ml of supernatant was transferred to a new tube, and 50 µl of protein-A agarose (Boehringer Mannheim, Indianapolis, Ind.) was added. The sample was incubated overnight with rotation, and then the beads were pelleted by centrifugation at 12,000 x g for 20 s. The supernatant was removed to a new tube and incubated with 20 µl of anti-AplA antibody with rotation for 3 h. After this period, 20 µl of protein A-agarose was added, and rotation was continued for another 3 h. The beads were then collected by centrifugation at 12,000 x g for 20 s and washed four times with 1 ml of wash buffer (50 mM Tris-HCl [pH 7.5], 500 mM NaCl, 0.5% [wt/vol] Triton X-100). Beads were resuspended in 35 µl of 2x SDS loading buffer, boiled for 3 min, and centrifuged for 1 min at 15,000 x g, and 30 µl of supernatant was separated by SDS-PAGE (34).

Proteins were separated by SDS-PAGE in 20% polyacrylamide gels. For immunoblot and zinc blot analyses, proteins were transblotted to PVDF (polyvinylidene difluoride) membranes (Bio-Rad Laboratories). To detect proteins carrying bilin adducts, the blots were incubated in 1.3 M zinc acetate for 1 h at room temperature (35) and then imaged with a Molecular Dynamics Storm 860 imager, using the red laser and a photomultiplier tube setting of 1,000 V to monitor fluorescence from bilin adducts. For antibody probes, membranes were incubated with 5% (wt/vol) nonfat dry milk in Tris-buffered saline (20 mM Tris-HCl [pH 7.5], 140 mM NaCl) for 1 h at room temperature, followed by probing with affinity-purified anti-AplA or anti-AP antibody. Primary antibody interactions were detected with IRDye800-conjugated affinity-purified anti-rabbit immunoglobulin G (Rockland, Gilbertsville, Pa.) by using an Odyssey imaging system (Licor, Lincoln, Nebr.).

Site-specific inactivation of aplA. We designed primers APLA1 (5' CGGGATCCATCGCAAAGCTATTGAACGGATGA 3') and APLA2 (5' CGGAGCTCGCTCAAGGTCGTCAACTCGA 3') to amplify a 3.9-kb genomic fragment containing aplA. These primers, including BamHI and SacI restriction sites (underlined), respectively, were used in a standard PCR using template DNA isolated from wild-type F. diplosiphon cells. The PCR fragment was digested with BamHI and SacI and subcloned into the similarly digested pRL278 suicide vector (8). The resulting plasmid was digested at the PvuI site located within aplA and blunted using the Klenow fragment. A 1.6-kb blunt-ended fragment containing nptI, encoding kanamycin resistance (13), was then ligated into the vector. The resulting plasmid, pRL278-AplAKan, was used for transformation of wild-type and FdBk cells as previously described (33). Double recombinants were selected essentially as described by Cai and Wolk (10).

AplA overexpression in F. diplosiphon. Plasmid pB32A was created by cloning the HindIII fragment of F. diplosiphon genomic DNA containing aplA into the same site within pGEM-7Zf(+) (Promega, Madison, Wis.). pB32A was cut with BsmAI and end filled, and a 1.1-kbp fragment containing aplA was isolated and cloned (in the same orientation as the kanamycin resistance gene) into pPL2.7 that had been BamHI cut and end filled to create pPL2.7-AplA. The plasmid pPL2.7-AplA was used to transform wild-type and FdBk cells as previously described (33). Phycobiliproteins were extracted from AplA overexpression and parent lines and quantified as previously described (4). Total soluble protein from overexpression and parental lines was isolated as described above. One hundred micrograms of total soluble protein was separated by SDS-PAGE, blotted onto a PVDF membrane, and probed with anti-AplA antibodies as described above to determine the degree of AplA overexpression.

Nucleotide sequence accession number. The DNA sequence of aplA from F. diplosiphon has been deposited in the GenBank database under accession number AY729019.


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RESULTS
 
Bioinformatics analysis of aplA and related genes. The aplA gene was flanked by three previously identified F. diplosiphon genes. Two of these, rcaE and rcaF, were located 3' of aplA and positioned in the opposite orientation (Fig. 1A). RcaE and RcaF were previously identified as sensor kinase and response regulator class proteins that are part of a regulatory pathway controlling complementary chromatic adaptation (CCA), an acclimation process that alters PBS composition in response to changes in light color, particularly in red light (RL) and GL (30-32, 48). Also, one of the two genes 5' of aplA, named cotB, was critical for normal expression of PE and the PE linkers during GL growth (Fig. 1A) (4).



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  		FIG. 1. Cyanobacterial genomic DNA regions containing aplA genes. For the F. diplosiphon (A) and N. punctiforme (B) genomes, the locations of aplA genes, of cotB genes, and of known regulatory genes rcaE, rcaF, and cpeR are shown by black, hatched, and gray arrows, respectively.

Comparisons of AplA to the GenBank database revealed similarity to a range of AP subunits from a number of species. The best match was to the AP {gamma} chain from the red alga Porphyra purpurea, which shared 30% identity and 53% similarity over 147 aa. AplA had similar relatedness to unorthodox AP subunits from F. diplosiphon, with 30% identity and 49% similarity over 150 aa to {alpha}APB and 28% identity and 47% similarity over 151 aa to ApcA2. Similarity to orthodox AP {alpha} and AP ß peptides was weaker, with 25% identity for both over stretches of 150 and 95 aa, respectively. Sequence alignment of AplA with AP class proteins from several cyanobacteria is provided in Fig. 2A.



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  		FIG. 2. Alignment of the deduced sequence of the AplA protein from F. diplosiphon with AP {alpha} (ApcA) and AP ß (ApcB) subunits (A) and with Apl proteins from several cyanobacterial species (B). Abbreviations: Fd, F. diplosiphon; Np, N. punctiforme; Te, T. erythraeum strain IMS101. Black backgrounds indicate 100% identical residues, while gray backgrounds indicate residues with at least 50% identity. Closed triangles indicate eight residues conserved in all phycobiliproteins; closed circles indicate eight residues primarily conserved in AP {alpha} proteins.

Two hypothetical proteins in the database matched AplA much more closely than AP subunit proteins (Fig. 2B). These were encoded in the genomes of the cyanobacteria N. punctiforme and T. erythraeum IMS101 which, like F. diplosiphon, synthesize PE. AplA was 66% identical and 75% similar over 172 aa to the N. punctiforme protein (AplANp) and 57% identical and 69% similar to the T. erythraeum protein (AplATe). The predicted lengths of AplANp and AplATe were 193 and 192 aa, respectively, which made them virtually identical in length to AplA, which had a predicted length of 193 aa, a theoretical pI of 5.38, and a calculated molecular weight of 21,964 (starting from the first valine). These three proteins also had conserved N-terminal extensions of approximately 28 aa that were absent in AP {alpha} and ß subunits, which are typically only 160 aa long (Fig. 2). Protein family database analysis of this extension did not detect any related region, and helix-turn-helix analysis did not identify a helix-turn-helix motif (data not shown).

Structural and functional studies of many phycobiliproteins, including AP {alpha} and ß subunits, have identified eight residues that are universally conserved in phycobiliproteins (Fig. 2A) (7). Based on F. diplosiphon ApcA1 protein numbering (Fig. 2A), these residues are D12, C81, R83, D84, R90, Y94, G99, and G111. The Apl class proteins were examined for conservation of these residues (Table 1). The first six (D12, C81, R83, D84, R90, and Y94) are completely conserved in all three members of the Apl family. Three of these (C81, R83, and D84) are critical for chromophore attachment (C81) or alignment (R83 and D84), while charge interactions among the other three (D12, R90, and Y94) are important for {alpha}-ß subunit pairing. The final two residues, G99 and G111, which appear to be necessary for achieving the proper tertiary structure of each individual subunit (3), are not conserved at five out of six positions within the three Apl family members (Fig. 2B).


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  		TABLE 1. Presence of conserved phycobiliprotein residues in Ap1Aa

There are also a number of residues that are primarily conserved within the AP {alpha} subfamily (Fig. 2A) (7). In F. diplosiphon ApcA1, these are E14, V66, M77 (L77 in the ApcA1 numbering convention), T78, T80, I107, Y116, and P122 (Table 1). The first six are important in {alpha}-ß subunit or {alpha} subunit-linker interactions, and two of these, M77 and T80, are also in contact with the chromophore. The final two (Y116 and P122) have important chromophore contacts. There is virtually no conservation of the first six residues in members of the Apl family, while the final two residues are 100% conserved in all three Apl proteins (Table 1).

Based on its position in the F. diplosiphon genome, aplA is not transcribed with any other genes and is not located within 5 kbp of any open reading frame encoding a protein with measurable sequence similarity to AP subunits or any other phycobiliproteins (Fig. 1A and data not shown). The absence of a neighboring AP gene and the proximity of aplA to three genes encoding proteins that regulate phycobiliprotein gene expression led us to examine the region of the genome encoding the most closely related protein, AplANp. In N. punctiforme, the gene encoding AplANp is located next to and divergently transcribed from a gene encoding a CotB homolog (4) (Fig. 1B). Thus, the arrangements of aplA and cotB homologs are nearly identical in F. diplosiphon and N. punctiforme, closely related filamentous cyanobacteria that both undergo chromatic adaptation (47). The only difference is the presence of an additional open reading frame between aplA and cotB in the F. diplosiphon genome. Downstream of the N. punctiforme aplA gene and transcribed in the same direction are homologs of cpeE, cpeS, cpeT, and cpeR (12) (GenBank accession no. AAAY02000033). In F. diplosiphon, cpeE encodes a PE linker peptide (16, 22) and is also located upstream of cpeS, cpeT, and cpeR (12, 42). The roles of CpeS and CpeT are unknown, but CpeR is required for cpeBA operon expression (12, 42). Thus, as in F. diplosiphon, the N. punctiforme aplA gene is not near a partner AP-encoding gene and is flanked by genes encoding regulators of phycobiliprotein gene expression.

AplA was compared to other proteins with sequence similarity to phycobiliproteins by use of the ProtML phylogenetic analysis program. AplA, AplANp, and AplATe were located in a single cluster distinct in primary sequence from the related groups containing the AP {alpha} and ß subunits (Fig. 3). The same results were obtained when this analysis was performed either with or without inclusion of the approximately 28-aa N-terminal extensions of AplA, AplANp, and AplATe (data not shown). AP {alpha} subunits clustered into a single group, while AP ß subunits fell into two groups, the smaller of which was more closely related to the AP {alpha} cluster. The bootstrap values obtained provided strong support for the tree shown, which supports the likelihood that AP subunits and the distinct Apl proteins arose from gene divergence following duplication of a common ancestor.



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  		FIG. 3. Phylogenetic analysis of cyanobacterial AplA and Apc proteins. Abbreviations for species: Acyl, Anabaena cylindrica; Ccal, Cyanidium caldarium; Fdip, F. diplosiphon; Nos7120, Nostoc sp. strain PCC 7120; Npun, N. punctiforme; Spla, Spirulina platensis; Syn6803, Synechocystis sp. strain PCC 6803; Syn7002, Synechococcus sp. strain PCC 7002; Telo, Thermosynechococcus elongatus BP-1; Tery, Trichodesmium erythraeum IMS101. Bootstrap values (in percentages), if greater than 50, are provided. A scale bar representing the number of substitutions per 100 sites is provided.

Three-dimensional structural modeling resulted in a predicted structure for AplA (Table 2) that, based on Fischer's fold recognition algorithm, was significantly similar (i.e., with a Z score of ≥7) (18) to a number of phycobiliproteins, including the {alpha} and ß subunits of AP and PC. In agreement with this, the FUGUE program predicted the AplA structure to be highly similar to the phycobiliprotein-containing "phc" family profile in the HOMSTRAD database (http://www-cryst.bioc.cam.ac.uk//homstrad/) (40) with a Z score of 26.78, where a Z score of ≥6 corresponds to a 99% confidence in the structure homology prediction.


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  		TABLE 2. Homology recognition results

FUGUE results were also used for structural model generation (43), using RasMac version 2.5. The AplA model was nearly indistinguishable from that of AP {alpha} (PDB entry 1alla) and contained helices with lengths and positions similar to those found for both AP subunits (data not shown). The most notable difference was a substitution of the basic R residue in AplA for the neutral M residue (M77, or L77 in the ApcA1 numbering convention) near the junction of helices E and G. Although this and other sequence divergences (Fig. 2A) have no major impact on the overall tertiary structure of AplA compared to the impact on that of AP, these changes are likely to have some effect on dimerization interactions and possibly on chromophore conformation.

AplA is expressed but not present in PBS. The evidence presented above suggested that although AplA is a member of the phycobiliprotein superfamily, it might not be incorporated into PBS in F. diplosiphon. AplA-specific antibodies were generated and used to determine whether AplA was present in PBS and differentially expressed in RL and GL. While control antibodies identified PBS components both in the supernatant fraction of whole-cell extracts and in purified PBS preparations, anti-AplA antibodies detected AplA at equal levels in RL and GL in the supernatant fraction of whole-cell extracts but not in purified PBS (Fig. 4, upper panels). AplA was also not detected in the insoluble protein fraction under either light condition (data not shown). SDS-PAGE and Coomassie blue staining of aliquots of the same protein samples demonstrated the degree of purification of the PBS components and the fact that the lack of AplA in the PBS was not due to any significant differences in the amount of PBS components present in each lane (Fig. 4, lower panels). Although AplA was predicted to be 193 aa in length, it migrated slightly faster than the major AP {alpha} and ß subunits, which are each 161 aa in length (Fig. 4, lower panels). The reason for this is not presently clear, as the strong sequence similarity of the N-terminal extension among Apl family members makes it likely that the predicted start sites, and thus lengths, of the Apl proteins are approximately correct. Attempts to sequence the N terminus of the protein were unsuccessful and thus did not provide additional insight into the size anomaly observed for AplA on SDS-PAGE gels.



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  		FIG. 4. AplA is not found in purified PBS of F. diplosiphon. Immunoblots show that anti-AP antibodies bound proteins in both whole-cell soluble (sol) and purified PBS (PBS) extracts (upper left panel), while anti-AplA antibodies bound protein in whole-cell soluble extracts but not protein in purified PBS (upper right panel). For each extract, samples were obtained from cells grown in RL and GL. Lower panels are Coomassie blue-stained gels containing equivalent aliquots of the same proteins used in the upper panels. Molecular weight standards (in thousands) are provided to the left of the gels, and major PBS components are labeled.

AplA covalently attaches a chromophore in vivo. AplA contained the chromophore attachment cysteine and many other conserved residues that contact the chromophore (Fig. 2A and Table 1). However, the absence of this protein in PBS (Fig. 4, upper right panel) raised the question of whether AplA was a photoreceptor, i.e., contained a covalently attached chromophore in vivo. Anti-AplA antibodies were used to immunoprecipitate AplA from the supernatant fraction of F. diplosiphon whole-cell protein extracts. A protein of the size previously observed for AplA that showed zinc-dependent fluorescence was immunoprecipitated specifically with the AplA antibodies (Fig. 5), indicating the presence of a linear tetrapyrrole chromophore covalently attached to this protein (35). The high degree of binding specificity of our affinity-purified anti-AplA antibodies (Fig. 4, upper right panel) makes it unlikely that the immunoprecipitated protein is not AplA. Control immunoprecipitations lacking anti-AplA antibodies failed to precipitate any protein, and zinc-dependent fluorescence was not detectable (Fig. 5). Furthermore, Western analysis of the immunoprecipitations with anti-AP antibodies did not detect any cross-contamination of AP in the samples (data not shown).



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  		FIG. 5. AplA has a covalently attached chromophore in vivo. Immunoblot analysis (Ab) was performed with anti-AplA antibodies after immunoprecipitation using whole-cell soluble protein extracts from wild-type F. diplosiphon either with the same antibodies (Anti-AplA, left) or without a primary antibody (control, right). The same membranes were subjected to zinc blot analysis (zinc) to monitor chromophore fluorescence (34).

Inactivation of aplA in F. diplosiphon has no obvious effect on CCA. Plasmid pRL278 (8), a sacB-containing homologous recombination plasmid, was utilized to inactivate aplA in both wild-type and FdBk (rcaE-null) mutant F. diplosiphon cells. Insertional mutagenesis of aplA was confirmed by PCR analysis (Fig. 6, left panel). The wild-type inactivation line did not accumulate detectable amounts of AplA protein, as determined by immunoblot analysis (Fig. 6, right panel). Whole-cell spectral analyses of the CCA responses in the aplA mutant lines did not reveal significant differences between phycobiliprotein accumulation patterns and those of parent cell lines (data not shown).



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  		FIG. 6. Isolation of aplA mutants via allelic exchange. (Left) PCR amplification of the aplA region in the wild type (wt), the FdBk (rcaE-null) mutant, and aplA allelic-exchange mutants ({Delta}aplA). Numbers to the left represent sizes in kilobases. (Right) Coomassie blue-stained (left two lanes) and anti-AplA Western blot (right two lanes) results for the wild type (wt) and the aplA mutant ({Delta}aplA). Numbers to the left indicate sizes in kilodaltons.

AplA overexpression influences PBS protein and transcript expression in F. diplosiphon. To further determine the role of AplA in F. diplosiphon, plasmid pPL2.7-AplA was used to overexpress AplA in both wild-type and FdBk mutant F. diplosiphon cells (Fig. 7). Phycobiliprotein levels were quantified in nontransformed and overexpression lines (Table 3). AplA overexpression in wild-type cells resulted in increased PC and PE levels in both RL and GL relative to levels in wild-type cells without AplA overexpression, and cpcB2A2 RNA levels were higher in GL in the wild-type overexpression lines, while cpeBA RNA levels were lower in RL in the wild-type overexpression lines. However, overexpression of AplA in an FdBk mutant background, which disabled the Rca system, gave quite different results. Relative to the nontransformed lines, FdBk mutant cells overexpressing AplA showed altered phenotypes only during growth in GL, where PE levels were only 65% of those measured in the FdBk mutant and PC levels were 124% higher. These results were mirrored in RNA accumulation patterns during growth in GL, where RNA levels in the overexpressing lines were just 38% (for cpeBA) and 71% (for cpeCDE) of those measured in FdBk mutant cells, while cpcB2A2 RNA levels were 151% of those in FdBk mutant cells (Table 3).



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  		FIG. 7. Overexpression of AplA in F. diplosiphon wild-type and FdBk mutant cells. Immunoblot analysis was performed with anti-AplA antibodies using whole-cell soluble protein extracts from the wild type (wt), the FdBk mutant, and the corresponding AplA overexpression (AOX) F. diplosiphon cell lines.


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  		TABLE 3. Phycobiliprotein ratios and RNA levels in AplA overexpression lines


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DISCUSSION
 
AplA is absent in purified PBS. AplA was undetectable in purified PBS preparations or insoluble protein fractions. This finding sets AplA apart from previously described phycobiliproteins, which have all been found within PBS (6, 23). AplA was present in the soluble portion of F. diplosiphon protein extracts (Fig. 4), which was prepared without detergent and subjected to high-speed centrifugation to reduce the possibility that AplA was stripped from complexes or membranes during extraction or that the soluble fraction was contaminated with membrane material. Although these data strongly suggest that AplA is not present within the PBS and has a novel cellular function, it is possible that AplA was weakly associated with PBS and lost during the PBS purification procedure. If so, the sequence similarity of AplA to AP subunits suggests that it would most likely be associated with the core, as has been found for {alpha}APB and ß18. However, these PBS core components have been shown to copurify with PBS, unlike AplA (21, 24, 36-38, 46, 51). Thus, any association between AplA and PBS that would be weak enough to lead to its loss during PBS purification would have to be quite novel, and therefore we consider this possibility unlikely.

Sequence analyses suggest a novel structure and function for Apl proteins. Through analysis of residues conserved between Apl and AP {alpha} proteins, three residues involved in chromophore attachment and coordination (C81, R83, and D84) as well as two residues with chromophore contacts (Y116 and P122) that are conserved in all AP {alpha} subunits were also identified in Apl family members (Fig. 2 and Table 1). Thus, it appears that most of the common features of chromophore binding and association are shared between the AP {alpha} and the Apl families, which is supported by our finding that AplA binds a chromophore in vivo (Fig. 5).

In addition, Apl proteins contain the conserved residues involved in charge interactions between {alpha} and ß subunits (D12, R90, and Y94) but mostly lack the six AP {alpha}-specific residues that are important for subunit-subunit and subunit-linker interactions (E14, V66, M77 or L77, T78, T80, and I107), suggesting that Apl proteins dimerize with proteins other than AP ß. This hypothesis is supported by five additional pieces of evidence. First, with extended N termini, the conceptually translated Apl family members are larger than AP proteins (Fig. 2), which would make it difficult for them to properly pack into PBS in the traditional {alpha}/ß dimer structure. Second, two residues (G99 and G111) conserved in all phycobiliproteins and necessary for proper individual subunit conformation are not conserved in Apl proteins (Fig. 2A and Table 1), suggesting that there is at least some variation in the tertiary structure of these proteins relative to that of previously described phycobiliproteins. Third, no AP partner genes exist near any of the three apl genes (Fig. 1 and data not shown). Fourth, no AP subunits were detected in anti-AP antibody probings of AplA immunoprecipitation experiments (data not shown). Finally, AplA is not found within the PBS (Fig. 4).

The in vivo covalent attachment of a bilin chromophore to AplA demonstrates that it is a photoreceptor, but it is not yet clear whether its role is photosensory. There have been previous suggestions that phycobiliproteins have light-sensing roles. The phycoerythrocyanin {alpha} subunit of Mastigocladus laminosus shows photoreversible photochemistry and has been suggested as having a possible role in chromatic adaptation or another form of photomorphogenesis (53). In addition, the discovery of PE in a low-light-adapted strain of the marine photosynthetic prokaryote Prochlorococcus marinus (26) and subsequent identification of a cpeB-like gene, but not cpeA, in the high-light-adapted Prochlorococcus strain MED4 led to the suggestion that the cpeB gene product may serve a light-sensory role (27), a proposal supported by the fact that proteins encoded by cpeB-like genes are highly conserved in many high-light-adapted Prochlorococcus strains (45).

The clustering of the apl genes with light-responsive regulatory components (Fig. 1), along with the lack of AplA in purified PBS (Fig. 4), raises the possibility that Apl proteins have photoresponsive regulatory, rather than energy collection, roles. In this regard, whole-cell spectral analyses of aplA-null mutants demonstrated that AplA does not play a major role in controlling CCA. However, overexpression of AplA in an FdBk mutant background during growth in GL did result in decreased levels of both PE protein and cpeBA and cpeCDE RNA and increased levels of PC protein and cpcB2A2 RNA (Table 3). These data suggest that AplA may play a negative role in regulating cpeBA and cpeCDE expression and a positive role in controlling cpcB2A2 expression during growth in GL, but whether or not this role is photosensory remains to be determined. If Apl proteins play a sensory role, they would be expected to require a heterologous dimerization partner in order to function, since they lack an obvious signal output domain. Such a dimerization partner(s), if any, is presently unknown. In fact, the sequence divergence between Apl family members (Fig. 2B) would appear to be sufficient to accommodate diversity in both function and partnering.


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ACKNOWLEDGMENTS
 
This research was supported by an NSF postdoctoral fellowship in microbial biology to B.L.M. and NSF grants MCB 0084297 and 0416797 to D.M.K.

We thank Alex Glazer, Donald Bryant, and Wendy Schluchter for the gift of anti-allophycocyanin antibodies, Peter Wolk for providing homologous recombination plasmid pRL278, and Danny Rice and Jeffrey D. Palmer for assistance with phylogenetic analyses.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biology, Indiana University, 1001 E. Third St., Bloomington, IN 47405. Phone: (812) 856-4715. Fax: (812) 855-6705. E-mail: dkehoe{at}bio.indiana.edu. Back


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REFERENCES
 
    1
  1. Adachi, J., and M. Hasegawa. 1996. ProtML 2.3b3: maximum likelihood inference of protein phylogeny. Institute of Statistical Mathematics, Tokyo, Japan.
    2. 2
  2. 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]
    3. 3
  3. Apt, K. E., J. L. Collier, and A. R. Grossman. 1995. Evolution of the phycobiliproteins. J. Mol. Biol. 248:79-96.[CrossRef][Medline]
    4. 4
  4. Balabas, B. E., B. L. Montgomery, L. E. Ong, and D. M. Kehoe. 2003. CotB is essential for complete activation of green light-induced genes during complementary chromatic adaptation in Fremyella diplosiphon. Mol. Microbiol. 50:781-793.[CrossRef][Medline]
    5. 5
  5. Bateman, A., E. Birney, L. Cerruti, R. Durbin, L. Etwiller, S. R. Eddy, S. Griffiths-Jones, K. L. Howe, M. Marshall, and E. L. L. Sonnhammer. 2002. The Pfam protein families database. Nucleic Acids Res. 30:276-280.[Abstract/Free Full Text]
    6. 6
  6. Betz, M. 1997. One century of protein crystallography: the phycobiliproteins. Biol. Chem. 378:167-176.[Medline]
    7. 7
  7. Bickel, P. J., K. J. Kechris, P. C. Spector, G. J. Wedemayer, and A. N. Glazer. 2002. Finding important sites in protein sequences. Proc. Natl. Acad. Sci. USA 99:14764-14771.[Abstract/Free Full Text]
    8. 8
  8. Black, T. A., Y. P. Cai, and C. P. Wolk. 1993. Spatial expression and autoregulation of hetR, a gene involved in the control of heterocyst development in Anabaena. Mol. Microbiol. 9:77-84.[Medline]
    9. 9
  9. Brejc, K., R. Ficner, R. Huber, and S. Steinbacher. 1995. Isolation, crystallization, crystal-structure analysis and refinement of allophycocyanin from the cyanobacterium Spirulina platensis at 2.3 Å resolution. J. Mol. Biol. 249:424-440.[CrossRef][Medline]
    10. 10
  10. Cai, Y., and C. P. Wolk. 1990. Use of a conditionally lethal gene in Anabaena sp. strain PCC 7120 to select for double recombinants and to entrap insertion sequences. J. Bacteriol. 172:3138-3145.[Abstract/Free Full Text]
    11. 11
  11. Casey, E. S., D. M. Kehoe, and A. R. Grossman. 1997. Suppression of mutants aberrant in light intensity responses of complementary chromatic adaptation. J. Bacteriol. 179:4599-4606.[Abstract/Free Full Text]
    12. 12
  12. Cobley, J. G., A. C. Clark, S. Weerasurya, F. A. Queseda, J. Y. Xiao, N. Bandrapali, I. D'Silva, M. Thounaojam, J. F. Oda, T. Sumiyoshi, and M. H. Chu. 2002. CpeR is an activator required for expression of the phycoerythrin operon (cpeBA) in the cyanobacterium Fremyella diplosiphon and is encoded in the phycoerythrin linker-polypeptide operon (cpeCDESTR). Mol. Microbiol. 44:1517-1531.[CrossRef][Medline]
    13. 13
  13. Cobley, J. G., and R. D. Miranda. 1993. Construction of shuttle plasmids which can be efficiently mobilized from Escherichia coli into the chromatically adapting cyanobacterium, Fremyella diplosiphon. Plasmid 30:90-105.[CrossRef][Medline]
    14. 14
  14. Collier, J. L., and A. R. Grossman. 1992. Chlorosis induced by nutrient deprivation in Synechococcus sp. strain PCC 7942: not all bleaching is the same. J. Bacteriol. 174:4718-4726.[Abstract/Free Full Text]
    15. 15
  15. Dodd, I. B., and J. B. Egan. 1990. Improved detection of helix-turn-helix DNA-binding motifs in protein sequences. Nucleic Acids Res. 18:5019-5026.[Abstract/Free Full Text]
    16. 16
  16. Federspiel, N. A., and L. Scott. 1992. Characterization of a light-regulated gene encoding a new phycoerythrin-associated linker protein from the cyanobacterium Fremyella diplosiphon. J. Bacteriol. 174:5994-5998.[Abstract/Free Full Text]
    17. 17
  17. Fischer, D. 2000. Hybrid fold recognition: combining sequence derived properties with evolutionary information, p. 119-130. In Proceedings of the Pacific Symposium on Biocomputing 2000. World Scientific, Honolulu, Hawaii.
    18. 18
  18. Fischer, D., and D. Eisenberg. 1996. Protein fold recognition using sequence-derived predictions. Protein Sci. 5:947-955.[Medline]
    19. 19
  19. Gantt, E. 1981. Phycobilisomes. Annu. Rev. Plant Physiol. 32:327-347.
    20. 20
  20. Gindt, Y. M., J. H. Zhou, D. A. Bryant, and K. Sauer. 1992. Core mutations of Synechococcus sp. PCC 7002 phycobilisomes: a spectroscopic study. J. Photochem. Photobiol. B 15:75-89.[CrossRef][Medline]
    21. 21
  21. Gingrich, J. C., D. J. Lundell, and A. N. Glazer. 1983. Core substructure in cyanobacterial phycobilisomes. J. Cell. Biochem. 22:1-14.[CrossRef][Medline]
    22. 22
  22. Glauser, M., W. A. Sidler, K. W. Graham, D. A. Bryant, G. Frank, E. Wehrli, and H. Zuber. 1992. Three C-phycoerythrin-associated linker polypeptides in the phycobilisome of green-light-grown Calothrix sp. PCC 7601 (cyanobacteria). FEBS Lett. 297:19-23.[CrossRef][Medline]
    23. 23
  23. Glazer, A. N. 1984. Phycobilisome. A macromolecular complex optimized for light energy transfer. Biochim. Biophys. Acta 768:29-51.
    24. 24
  24. Glazer, A. N., and D. A. Bryant. 1975. Allophycocyanin B ({lambda}max 671, 618 nm). A new cyanobacterial phycobiliprotein. Arch. Microbiol. 104:15-22.[CrossRef][Medline]
    25. 25
  25. Grossman, A. R., D. Bhaya, K. E. Apt, and D. M. Kehoe. 1995. Light-harvesting complexes in oxygenic photosynthesis: diversity, control, and evolution. Annu. Rev. Genet. 291:231-288.[CrossRef]
    26. 26
  26. Hess, W. R., F. Partensky, G. W. M. van der Staay, J. M. Garcia-Fernandez, T. Börner, and D. Vaulot. 1996. Coexistence of phycoerythrin and a chlorophyll a/b antenna in a marine prokaryote. Proc. Natl. Acad. Sci. USA 93:11126-11130.[Abstract/Free Full Text]
    27. 27
  27. Hess, W. R., G. Rocap, C. S. Ting, F. Larimer, S. Stilwagen, J. Lamerdin, and S. W. Chisholm. 2001. The photosynthetic apparatus of Prochlorococcus: insights through comparative genomics. Photosynth. Res. 70:53-71.
    28. 28
  28. Houmard, J., V. Capuano, T. Coursin, and N. Tandeau de Marsac. 1988. Genes encoding core components of the phycobilisome in the cyanobacterium Calothrix sp. strain PCC 7601: occurrence of a multigene family. J. Bacteriol. 170:5512-5521.[Abstract/Free Full Text]
    29. 29
  29. Houmard, J., V. Capuano, T. Coursin, and N. Tandeau de Marsac. 1988. Isolation and molecular characterization of the gene encoding allophycocyanin B, a terminal energy acceptor in cyanobacterial phycobilisomes. Mol. Microbiol. 2:101-107.[CrossRef][Medline]
    30. 30
  30. Kehoe, D. M., and A. R. Grossman. 1994. Complementary chromatic adaptation: photoperception to gene regulation. Semin. Cell Biol. 5:303-313.[Medline]
    31. 31
  31. Kehoe, D. M., and A. R. Grossman. 1997. New classes of mutants in complementary chromatic adaptation provide evidence for a novel four-step phosphorelay system. J. Bacteriol. 179:3914-3921.[Abstract/Free Full Text]
    32. 32
  32. Kehoe, D. M., and A. R. Grossman. 1996. Similarity of a chromatic adaptation sensor to phytochrome and ethylene receptors. Science 273:1409-1412.[Abstract]
    33. 33
  33. Kehoe, D. M., and A. R. Grossman. 1998. Use of molecular genetics to investigate complementary chromatic adaptation: advances in transformation and complementation. Methods Enzymol. 297:279-290.[CrossRef]
    34. 34
  34. Laemmli, U. K. 1970. Cleavage of structural protein during the assembly of the head of bacteriophage T4. Nature 227:680-685.[CrossRef][Medline]
    35. 35
  35. Li, L., and J. C. Lagarias. 1992. Phytochrome assembly. Defining chromophore structural requirements for covalent attachment and photoreversibility. J. Biol. Chem. 267:19204-19210.[Abstract/Free Full Text]
    36. 36
  36. Lundell, D. J., and A. N. Glazer. 1981. Allophycocyanin B. J. Biol. Chem. 256:12600-12606.[Abstract/Free Full Text]
    37. 37
  37. Lundell, D. J., and A. N. Glazer. 1983. Molecular architecture of a light-harvesting antenna. Core substructure in Synechococcus 6301 phycobilisomes: two new allophycocyanin and allophycocyanin B complexes. J. Biol. Chem. 258:902-908.[Abstract/Free Full Text]
    38. 38
  38. Lundell, D. J., and A. N. Glazer. 1983. Molecular architecture of a light-harvesting antenna. Structure of the 18 S core-rod subassembly of the Synechococcus 6301 phycobilisome. J. Biol. Chem. 258:894-901.[Abstract/Free Full Text]
    39. 39
  39. Maxson, P., K. Sauer, J. H. Zhou, D. A. Bryant, and A. N. Glazer. 1989. Spectroscopic studies of cyanobacterial phycobilisomes lacking core polypeptides. Biochim. Biophys. Acta 977:40-51.[Medline]
    40. 40
  40. Mizuguchi, K., C. M. Deane, T. L. Blundell, and J. P. Overington. 1998. HOMSTRAD: a database of protein structure alignments for homologous families. Protein Sci. 7:2469-2471.[Medline]
    41. 41
  41. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
    42. 42
  42. Seib, L. O., and D. M. Kehoe. 2002. A turquoise mutant genetically separates expression of genes encoding phycoerythrin and its associated linker peptides. J. Bacteriol. 184:962-970.[Abstract/Free Full Text]
    43. 43
  43. Shi, J. Y., T. L. Blundell, and K. Mizuguchi. 2001. FUGUE: sequence-structure homology recognition using environment-specific substitution tables and structure-dependent gap penalties. J. Mol. Biol. 310:243-257.[CrossRef][Medline]
    44. 44
  44. Sidler, W. A. 1994. Phycobilisome and phycobiliprotein structures, p. 139-216. In D. A. Bryant (ed.), The molecular biology of cyanobacteria. Kluwer, Dordrecht, The Netherlands.
    45. 45
  45. Steglich, C., A. F. Post, and W. R. Hess. 2003. Analysis of natural populations of Prochlorococcus spp. in the northern Red Sea using phycoerythrin gene sequences. Environ. Microbiol. 5:681-690.[CrossRef][Medline]
    46. 46
  46. Suter, F., P. Fuglistaller, D. J. Lundell, A. N. Glazer, and H. Zuber. 1987. Amino acid sequences of {alpha}-allophycocyanin B from Synechococcus 6301 and Mastigocladus laminosus. FEBS Lett. 217:279-282.[CrossRef]
    47. 47
  47. Tandeau de Marsac, N. 1977. Occurrence and nature of chromatic adaptation in cyanobacteria. J. Bacteriol. 130:82-91.[Abstract/Free Full Text]
    48. 48
  48. Tandeau de Marsac, N. 1983. Phycobilisomes and complementary chromatic adaptation in cyanobacteria. Bull. Inst. Pasteur 81:201-254.
    49. 49
  49. Tandeau de Marsac, N., and G. Cohen-Bazire. 1977. Molecular composition of cyanobacterial phycobilisomes. Proc. Natl. Acad. Sci. USA 74:1635-1639.[Abstract/Free Full Text]
    50. 50
  50. Thompson, J. D., D. G. Higgins, and T. J. Gibson. 1994. CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res. 22:4673-4680.[Abstract/Free Full Text]
    51. 51
  51. Yamanaka, G., D. J. Lundell, and A. N. Glazer. 1982. Molecular architecture of a light-harvesting antenna. Isolation and characterization of phycobilisome subassembly particles. J. Biol. Chem. 257:4077-4086.[Abstract/Free Full Text]
    52. 52
  52. Zhao, J., J. Zhou, and D. Bryant. 1992. Energy transfer process in phycobilisomes as deduced from analyses of mutants of Synechococcus sp. PCC 7002, p. 25-32. In N. Murata (ed.), Research in photosynthesis, vol. I. Kluwer, Dordrecht, The Netherlands.
    53. 53
  53. Zhao, K.-H., and H. Scheer. 1995. Type I and type II reversible photochemistry of phycoerythrocyanin {alpha}-subunit from Mastigocladus laminosus both involve Z, E isomerization of phycoviolobilin chromophore and are controlled by sulfhydryls in apoprotein. Biochim. Biophys. Acta Bioenergetics 1228:244-253.[CrossRef]

Journal of Bacteriology, November 2004, p. 7420-7428, Vol. 186, No. 21
0021-9193/04/$08.00+0     DOI: 10.1128/JB.186.21.7420-7428.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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