©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Assembly of the Photosystem II Oxygen-evolving Complex Is Inhibited in psbA Site-directed Mutants of Chlamydomonas reinhardtii
ASPARTATE 170 OF THE D1 POLYPEPTIDE (*)

(Received for publication, August 25, 1994; and in revised form, October 18, 1994)

Julian P. Whitelegge (1) Derrick Koo (1) Bruce A. Diner (2) Ibrahim Domian (1) Jeanne M. Erickson (1) (3)(§)

From the  (1)Department of Biology, University of California, Los Angeles, California 90024, (2)Central Research and Development Department, E.I. Du Pont de Nemours and Company, Wilmington, Delaware 19880-0173, and the (3)Molecular Biology Institute, University of California, Los Angeles, California 90024-1606

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Photosystem II catalyzes the photooxidation of water to molecular oxygen, providing electrons to the photosynthetic electron transfer chain. The D1 and D2 chloroplast-encoded reaction center polypeptides bind cofactors essential for Photosystem II function. Transformation of the chloroplast genome of the eukaryotic green alga Chlamydomonas reinhardtii has allowed us to engineer site-directed mutants in which aspartate residue 170 of D1 is replaced by histidine (D170H), asparagine (D170N), threonine (D170T), or proline (D170P). Mutants D170T and D170P are completely deficient in oxygen evolution, but retain normal (D170T) or 50% (D170P) levels of Photosystem II reaction centers. D170H and D170N accumulate wild-type levels of PSII centers, yet evolve oxygen at rates approximately 45% and 15% those of control cells, respectively. Kinetic analysis of chlorophyll fluorescence in the mutants reveals a specific defect in electron donation to the reaction center. Measurements of oxygen flash yields in D170H show, however, that those reaction centers capable of evolving oxygen function normally. We conclude that aspartate residue 170 of the D1 polypeptide plays a critical role in the initial binding of manganese as the functional chloroplast oxygen-evolving complex is assembled.


INTRODUCTION

In photosynthetic eukaryotes, the Photosystem II (PSII) reaction center mediates the photooxidation of water to molecular oxygen, providing electrons to the photosynthetic electron transfer chain and releasing protons into the lumen of the chloroplast thylakoid membrane. At least five nuclear-encoded and thirteen chloroplast-encoded PSII polypeptide subunits have been identified (for reviews, see Erickson and Rochaix(1992) and Ikeuchi(1992)). The function of PSII centers active on both the donor and acceptor sides, as outlined in Fig. 1and described in the legend, requires the interaction of the PSII polypeptides and their assembly with the lipid, metal, and ionic cofactors of PSII. At the heart of PSII structure and function is a heterodimer composed of two intrinsic membrane polypeptides, D1 and D2 (Nanba and Satoh, 1987; Marder et al., 1987; Webber et al., 1989; Tang et al., 1990). These two chloroplast-encoded polypeptides provide ligands for the reaction center chlorophyll P, pheophytin, and the primary and secondary quinone acceptor molecules of PSII, Q(A) and Q(B), respectively (Trebst, 1987; Svensson et al., 1990; Ruffle et al., 1991).


Figure 1: Schematic diagram of electron transfer events mediated by Photosystem II. Arrows indicate the linear transfer of electrons from water (H(2)O) to the second quinone acceptor molecule (Q(B)). The D1 and D2 polypeptides of the PSII reaction-center core provide ligands to redox active components on both the donor and acceptor sides of the PSII reaction center chlorophyll P, as indicated. Absorption of light energy (h) by the primary donor, P, leads to formation of the initial charge-separated state between P and a pheophytin a molecule (P-Pheo). Charge separation is stabilized by further transfer of the electron from pheophytin to the first and then second quinone acceptors, Q(A) and Q(B), respectively. This latter step is blocked by the herbicide diuron (DCMU) which binds competitively with quinone at the Q(B)-binding site of the D1 polypeptide. P is reduced by the secondary donor Z (Y(Z)), identified by Debus et al.(1988) and Metz et al.(1989) as tyrosine residue 161 of the D1 polypeptide (D1Y161). The tyrosine radical is reduced, in turn, by the functional oxygen-evolving complex (OEC) associated with the PSII core complex. According to the model of Joliot and Kok, with each photooxidation of the PSII reaction center, oxidizing equivalents are transferred one at a time to the OEC, advancing the so-called S-states of the OEC. The number associated with each S state (S(0)-S(4)) indicates the number of oxidizing equivalents stored (Kok et al., 1970; Joliot and Kok, 1975). Oxygen is released at the transient formation of the S(4) state restoring the S(0) state. The cycle through the S-s tates is thought to occur largely through oxidation of a tetranuclear manganese cluster at the heart of the OEC which supplies one electron for the reduction of P after each photooxidation of chlorophyll and accumulates the 4 oxidizing eq needed to split water. Current models predict that D1 and possibly D2 provide ligands to the manganese cluster.



Central to PSII water oxidation is a cluster of 4 manganese ions which advances in oxidation state as electrons are donated from it to the photooxidized PSII reaction center chlorophyll, P. When 4 oxidizing eq are accumulated in the manganese cluster, 1 molecule of oxygen is released and the cycle of manganese oxidation begins again (see Fig. 1 legend). The minimal, purified chloroplast PSII particle that retains maximal oxygen-evolving activity in vitro, in the absence of elevated levels of Ca or Cl (reviewed in Critchley (1985) and Andersson and Åkerlund(1987)), consists of the D1bulletD2 heterodimer, several additional intrinsic membrane polypeptides, and three extrinsic membrane polypeptides referred to as the oxygen evolution enhancer (OEE) (^1)polypeptides, OEE1, OEE2, and OEE3, of approximately 33, 23, and 17 kDa, respectively. The OEE polypeptides are localized to the lumenal side of PSII and participate with the intrinsic PSII polypeptides in the formation of a fully activated chloroplast oxygen-evolving complex (OEC) (reviewed in Andersson and Åkerlund(1987), Ghanotakis and Yocum (1990), Debus(1992), and Rutherford et al.(1992)). It is generally agreed that the OEC active site, containing Ca and 4 manganese atoms, requires 20 to 24 coordinating ligands provided mainly by oxygen, but with some nitrogen (DeRose et al., 1991). Histidine is now known to be a ligand (Tang et al., 1994), and additional coordination to manganese may be provided by carboxylate and amide moieties and perhaps tyrosine. Most evidence suggests that it is the D1 and, possibly, D2 polypeptides which coordinate manganese (reviewed in Diner et al.(1991), Debus (1992), and Vermaas and Pakrasi(1992)). Aspartate residue 170 (Asp-170) of the D1 polypeptide is found in the lumen only 9 amino acid residues away from D1 tyrosine 161 (Y) which directly accepts electrons from manganese (Debus et al., 1988; Metz et al., 1989). Asp-170 is conserved in the predicted D1 sequences of all 38 species of photosynthetic eukaryotes and cyanobacteria reviewed (Svensson et al., 1991). The role of aspartate 170 of the D1 polypeptide was recently examined in the cyanobacterium Synechocystis 6803 (Nixon and Diner, 1992; Boerner et al., 1992). These authors found that substitutions at this site abolished or reduced oxygen evolution in mutant cyanobacteria. Results of in vitro spectroscopic analysis of PSII particles isolated from mutant cyanobacteria suggest that D1 aspartate 170 plays an important role in the stable assembly of the manganese cluster of the cyanobacterial OEC.

Although photosynthesis in the prokaryotic cyanobacteria is similar in many respects to photosynthetic function in eukaryotic plants and algae, there are several features unique to the structure, function, regulation, and stability of the photosynthetic apparatus of the chloroplast. In eukaryotes, the photosystems are located in a thylakoid membrane within the chloroplast and, hence, totally isolated from the plasma membrane of the cell. Moreover, since the genes encoding polypeptides localized to the chloroplast thylakoid membrane are contained in two distinct genetic compartments as noted above, assembly of the chloroplast photosynthetic apparatus requires the coordinate expression of genes located in the nuclear genome and in the chloroplast genome, the latter of which is polyploid. This genetic complexity is in contrast to the prokaryotic cyanobacteria which lack intracellular organelles and which contain only one genome. Cyanobacteria also lack the highly ordered stacking of thylakoids into appressed and nonappressed membrane regions. There are substantial differences between cyanobacteria and chloroplast thylakoids with respect to the light-harvesting antennae polypeptides and pigments associated with the photosystems, the regulatory mechanisms affecting photosynthetic function (e.g. phosphorylation of the chloroplast reaction center polypeptides), and the ratio of PSII to PSI centers. There are also differences in PSII polypeptide composition. Cyanobacteria lack homologues for the OEE2 and OEE3 polypeptides (Stewart et al., 1985; Shen et al., 1992), and, although an OEE1 homologue exists, site-directed mutagenesis of the cyanobacterial OEE1 gene suggests that OEE1 is not essential for cyanobacterial PSII oxygen evolution in vivo (Bockholt et al., 1991; Mayes et al., 1991; Philbrick et al., 1991; Burnap et al., 1992). This is in marked contrast to the requirement for OEE1 and OEE2 in chloroplast oxygen evolution in vivo (Mayfield et al., 1987a, 1987b). Finally, differences in the interaction of PSII subunits are reflected by the fact that, in chloroplasts, loss of one subunit constituent has a more drastic consequence on the stability of the entire PSII complex (reviewed in Rochaix(1992)).

There has been until now little evidence as to whether specific changes in PSII polypeptide structure that affect PSII function in cyanobacteria would have a similar affect on the chloroplastic PSII. Given the above differences between photosynthesis in cyanobacteria and in the chloroplast of eukaryotes, we have undertaken studies to examine the relationship between PSII structure and donor-side function in a eukaryotic photosynthetic organism, C. reinhardtii. This unicellular green alga provides an excellent experimental system for a molecular and genetic approach to dissecting chloroplast PSII structure-function relationships (Rochaix and Erickson, 1988). DNA-mediated transformation of the nuclear (Debuchy et al., 1989; Fernandez et al., 1989; Mayfield and Kindle, 1990) and chloroplast (Boynton et al., 1988) genomes of this algae has been successful. Advantages of using Chlamydomonas to study PSII function include the much greater variable fluorescence compared to cyanobacteria, which allows for the rapid screening of algal colonies for mutant fluorescence phenotypes indicative of mutant PSII function, and the ability of Chlamydomonas to grow heterotrophically in the light or dark, which allows for easy maintenance of mutant strains deficient in photosynthesis. Here we report the site-directed mutagenesis of codon 170 of the chloroplast psbA gene encoding the D1 polypeptide of C. reinhardtii, the in vivo characterization of PSII function in algal transformants with amino acid substitutions at D1 residue 170, and our conclusions that residue 170 is critical for assembly and/or stability of manganese in the OEC of the chloroplast PSII.


EXPERIMENTAL PROCEDURES

Bacterial and Algal Strains

Escherichia coli strains C600 and DH5alpha (Sambrook et al., 1989) and MV1190 (Messing, 1983) were used as hosts for cloning and production of plasmid DNAs. The E. coli dutung strain CJ236 (Kunkel et al., 1987) was obtained from Bio-Rad and used for production of single-stranded phagemid DNA containing uracil. C. reinhardtii 2137 mt and 137y mt wild-type photosynthetic strains a nd the psbA deletion-mutant strain FuD7 mt (Bennoun et al., 1986) were used as hosts for chloroplast transformation. The 2137 and FuD7 strains are green in the dark, while the 137y strain is green only when grown in the light. Growth medium containing acetate (TAP) and minimal medium (HSM) were prepared as in Rochaix et al.(1988). Host and transformant algal cells used for phenotypic assays were grown in TAP liquid medium under low light (10- 12µmol m s) at 25 °C and harvested when cultures reached midlog stage. Photon flux densities were measured using a quantum radiometer (LI 185B, LiCor, Inc.).

Plasmid DNA Constructs and Site-directed Mutagenesis

The C. reinhardtii chloroplast restriction fragments contained in plasmids pRR, pRX, pRRX, and pXb1.8 are shown in Fig. 2. The EcoRI chloroplast restriction fragment R16 containing psbA exons 1-4 was cloned in the plasmid vector pBR328 (Erickson et al., 1984) and is here called pRR. The 1.8-kb XbaI fragment containing part of exon 2 and all of exon 3 was subcloned from pRR into the Bluescript KS phagemid vector (Stratagene) to produce plasmid pXb1.8. Mutant pXb1.8 plasmids were produced by site-directed mutagenesis as described below. A derivative of plasmid pRR which lacks the 1.8-kb XbaI fragment (pRR-DeltaXb1.8) was obtained after digesting pRR DNA with XbaI and religating. Mutant pRR plasmids were constructed by inserting the Xb1.8 fragment isolated from a mutant Xb1.8 plasmid into the XbaI site of pRR-DeltaXb1.8. pRX was produced by cloning the 5.7-kb EcoRI/XhoI subfragment of C. reinhardtii chloroplast DNA EcoRI restriction fragment R24 (Rochaix, 1978) into the EcoRI/XhoI sites of the Bluescript SK vector. The wild-type pRRX was constructed by inserting the wild-type R16 DNA fragment isolated from pRR into the single EcoRI site of pRX. Mutant pRRX plasmids were constructed using mutant pRR and wild-type pRX. Digestion of pRRX plasmids with XbaI was used to verify the correct orientation of psbA exons 1-4 with respect to exon 5. pPX261, used as a probe for RFLP screening of site-directed algal transformants, was obtained by inserting the 261-bp PstI/XbaI fragment isolated from pXb1.8 (see Fig. 3C) into the Bluescript SK vector. Conditions for ligation and bacterial transformation are as described in Sambrook et al.(1989). Plasmid DNAs used for transformation of algal cells were CsCl or column (Qiagen) purified from bacterial lysates.


Figure 2: Restriction map of the C. reinhardtii chloroplast psbA and rRNA genes and the mutations introduced at psbA codon 170. Upper: black boxes indicate the five exons of psbA (1-5) and the 16 S, 7 S, 3 S, 23 S, and 5 S rRNA genes. Hatched boxes indicate the four psbA introns and the intron in the 23 S rRNA gene. Bars above the restriction map show the relative locations of psbA DNA fragments cloned into recombinant plasmids pXb1.8, pRR, pRX, and pRRX. pCrBH4.8 contains a mutant 16 S rRNA gene which confers spectinomycin resistance to cells (Harris et al., 1989; Newman et al., 1990). Restriction sites for EcoRI (R), XhoI (X), XbaI (Xb), and BamHI (B) are indicated. Size of the bar is 1 kb. Lower: the wild-type DNA sequence of a portion of psbA exon 3 containing aspartate codon 170 is shown. The site-directed mutations indicated below the arrow were introduced into psbA in four different plasmid constructs. Indicated for each mutant codon is the corresponding amino acid residue substituted at position 170, the restriction site created by the mutation (far right), and the code used to label mutant constructs (far left).




Figure 3: Restriction fragment length polymorphism (RFLP) analysis of Chlamydomonas transformants. Analysis of DNA isolated from spectinomycin-resistant C. reinhardtii transformants shows the segregation of wild-type and mutant chloroplast DNA BsrI fragments containing a portion of psbA exon 3. DNA samples were prepared from 1.5-ml liquid cultures derived from 8 independent initial isolates (panel A) obtained after co-transformation of the WT2137 algal host with pCrBH4.8 and pRR-D170T plasmid DNAs (see Fig. 2) and from subsequent colony isolates (panel B) produced from the initial isolate shown in lane 3 of panel A. DNAs were subjected to BsrI restriction digestion and 1.5% agarose gel electrophoresis. Panels A and B show exposures obtained by autoradiography of Southern blots hybridized with a radiolabeled probe (261-bp fragment from pPXb261) indicated above the restriction map in panel C. A portion of psbA exon 3 is indicated by the black box. Restriction enzyme sites are given for BsrI (Bs), PstI (P), and XbaI (Xb). The BsrI site introduced by the D170T mutation in exon 3 is indicated with a star. Bars below the map show the DNA region contained in the wild-type (WT) and mutant (D170T) BsrI fragments of 303 bp and 240 bp, respectively.



Site-directed mutagenesis targeted to codon 170 of psbA was performed using the protocols described (Mutagene Kit, Bio-Rad) except that Bluescript KS (Stratagene) was used as the phagemid vector, R408 (Stratagene) was used as the M13 helper phage, and kanamycin was not added to superinfected cells. A synthetic 25-mer oligonucleotide homologous to the region of psbA surrounding codon 170 (sequence given in Fig. 2) but mutant at the third position of the codon (T) and degenerate at the first (C/A/T) and second (A/C) positions, was used to prime second strand DNA synthesis.

Chloroplast Transformation and Selection and Screening of Transformants

Algal host cells were harvested from liquid culture at midlog stage and 2 times 10^7 cells spread over the surface of a Nytran nylon membrane filter (0.45 µm, 82-mm diameter, Schleicher and Schull) placed on a TAP agar plate. Tungsten M10 particles (0.4-1.5 µm in diameter, now distributed by Bio-Rad) were obtained from Biolistics Inc., Ithaca, NY. 2.5 mg of M10 tungsten particles were coated, as otherwise described by Sanford et al.(1993), with 2.5 µg each of a psbA transforming plasmid construct and the mutant 16 S rRNA spectinomycin resistance plasmid p228 (Newman et al., 1990; received from K. Kindle as pCrBH4.8, see Fig. 2). Biolistic transformation was carried out with the PDS-1000/He (DuPont NEN) as instructed by the manufacturer, with the macrocarrier positioned 1 cm from the rupture disc (1100 p.s.i.) and 1 cm from the stopping screen.

Bombarded plates were incubated 16 h in the dark, after which filters containing cells were transferred to selective medium. TAP-SPEC agar (165 µg of spectinomycin dihydrochloride/ml) selected for spectinomycin-resistant cells. The spectinomycin-resistant control transformant lines WTsr (from transformation of 137y with p228) and FWTsr (from co-transformation of FuD7 with wild-type pRRX and p228) were isolated. An additional control strain, FWT, was produced by transformation of FuD7 with wild-type pRRX DNA followed by selection for growth on minimal medium (HSM) in the light. Initial colonies were picked into liquid medium and replated on selective medium, and secondary colony isolates were obtained. Primary and secondary isolates were subjected to colony hybridization and/or RFLP screening. All incubations during recovery, selection, screening, and routine maintenance of transformant cell lines were carried out in the dark.

Southern Blot Analysis and Colony Hybridization

Nucleic acids were isolated from 50 µl of algal cell pellet as described (Newman et al., 1990) except that the Pronase was omitted from the lysis buffer. DNAs were digested with restriction enzymes, separated by agarose gel electrophoresis, and transferred to a nylon membrane filter (Genescreen Plus, DuPont NEN) according to the manufacturer's instructions. Hybridization was carried out overnight at 65 °C as described (Church and Gilbert, 1984). Radiolabeled probes (BRL Random Primers DNA labeling system) were prepared using gel-purified DNA restriction fragments and [alphaP]dATP (3000 Ci/mmol, Amersham). Autoradiograms were obtained with X-Omat AR film (XAR5, Kodak).

Algal colonies grown on nylon membrane filters were screened by colony hybridization using a modification of the method developed for bacterial colonies (Grunstein and Hogness, 1975) as follows. Nylon filters were placed colony side up, for 2 min each treatment, onto 0.75-ml puddles of the following solutions, in sequence: 10% SDS (3 times 0.75 ml); 0.5 M NaOH, 1.5 M NaCl (3 times 0.75 ml); and 0.5 M Tris-HCl, pH 8.0, 1.5 M NaCl (3 times 0.75 ml). After each puddle treatment, excess liquid was removed from the filter using a Büchner funnel connected to a vacuum line. Filters were processed and hybridized as described above for Southern blot analysis.

DNA Sequence Analysis from DNA and RNA Templates

The universal and KS Bluescript primers (Stratagene) and custom primers homologous to regions of psbA intron 2 and intron 3 were used for DNA sequence analysis (Sequenase, U. S. Biochemical Corp.) of mutant plasmid DNAs. Algal RNA was isolated (Johanningmeier et al., 1987), and dideoxy sequencing reactions using [alpha-S]dATP (1000 Ci/mmol, Amersham) and 10 µg of RNA template were carried out as described (Fichot and Girard, 1990). A synthetic oligonucleotide homologous to exon 4 (5`-TCTGCTTGGAATACGATCAT-3`) was used to prime sequencing of the mutation site in exon 3 and was suitable for reading sequences through to exon 1. A synthetic oligonucleotide homologous to exon 5 (5`-GTTGTTTGAGCTAGAGTT-3`) was used to prime sequencing from the primer through exons 4 and 3. Oligonucleotides were synthesized on a Pharmacia Gene Assembler Plus and purified using a glass matrix (Mermaid, Bio 101 Inc.). Sequencing gels containing 7 M urea and 5% or 8% acrylamide (Long Ranger, AT Biochem. Inc.) were prepared and run according to the manufacturer's instructions.

In Situ Fluorescence Induction

Algal cells were grown in low light (6 µmol m s) on TAP agar plates. Fluorescence induction transients were recorded from cell patches using the Heinz-Walz pulse-amplitude-modulated fluorimeter (Schreiber, 1986). Actinic illumination (1500 µmol m s, white incandescent light) was controlled by a shutter (opening time < 2 ms), and transients were stored using a microcomputer running the Heinz-Walz DA100 program. Cells were dark-adapted for at least 10 min prior to assay at room temperature.

Quantification of PSII

PSII in whole cells was quantified by measuring the number of [^14C]diuron binding sites which were sensitive to competitive dissociation in the presence of 20 µM nonlabeled atrazine (Vermaas et al., 1990). [^14C]diuron (Amersham, 57 Ci mol) was diluted in assay buffer to seven different concentrations in the range 5-100 nM, and whole cells were added such that each assay tube contained 50 µg of chlorophyll/ml. Chlorophyll concentration was determined in 80% acetone (Arnon, 1949).

Oxygen Evolution

Rates of oxygen evolution were measured using an oxygen electrode (Hansatech) connected to a chart recorder. Liquid cultures were incubated in darkness for 2 min prior to illumination with saturating white light (500 µmol m s). Rates of oxygen evolution were 200 µmol of O(2)/mg of chlorophyll/h for 2137, 137y, and WTsr. The control transformants FWT and FWTsr displayed maximal rates of oxygen evolution of 100 µmol of O(2)/mg of chlorophyll/h. Rates listed in Table 1are shown as percentages of control rates of WTsr for all transformants where a wild-type host was used and FWTsr where FuD7 was used as host. The minimal detectable rate of oxygen evolution was 0.2 µmol of O(2)/mg of chlorophyll/h.



Relaxation of Chlorophyll Fluorescence after Saturating Light Flashes

The rates of relaxation of the chlorophyll fluorescence yield following actinic flashes were measured in whole cells using a flash detection spectrophotometer as described (Nixon and Diner, 1992). Algal cells were suspended in TAP medium at an A of 0.5. In contrast to previous assays using cyanobacteria (Nixon and Diner, 1992), the algal cells were not pretreated with benzoquinone and ferricyanide. A fresh sample was used for each set of delay times between the actinic and detecting flashes.

Oxygen Yields after Saturating Light Flashes and Determination of the Rate of Decay of the S(2) State

C. reinhardtii cells in 2 ml of TAP medium, adjusted to contain an additional 100 mM KCl, were sedimented onto the surface of an 11-mm bare platinum disk electrode. This electrode was surrounded by a 1-mm-thick Ag/AgCl ring electrode polarized 0.6 V positive with respect to the Pt electrode. The cells were dark-adapted 5-10 min and then subjected to a series (533 ms apart) of saturating light flashes supplied by a xenon strobe (Gen Rad, 1539A). The oxygen signals were measured from 1-51 ms after the actinic flash and captured by an AC-coupled digital oscilloscope (Nicolet). The signals were transferred to a computer (IBM Model 30) that integrated and tabulated the data. The computer also controlled the number and timing of the xenon flashes. The S(2) state lifetimes were determined as in Forbush et al.(1971) (see Fig. 9B legend).


Figure 9: Oxygen yields from Chlamydomonas cells following saturating light flashes. Panel A, relative yields of oxygen were measured following each of a series of saturating light flashes (533 ms apart) given to dark-adapted whole cells, as described under ``Experimental Procedures.'' The FWTsr(a) (control) and D170H(c) (transformant) cells were dark-adapted for 10 min prior to illumination. Panel B, lifetime of the S(2) state for these cells under the same conditions as for A. The cells were dark-adapted for 5 min and then given a single saturating light flash. Following this flash and at the time intervals indicated, a series of flashes were given 533 ms apart and the relative amplitude of oxygen was measured on the third overall flash. The relative concentration of the S(2) state is plotted as the difference, between the third overall flash yield for delay time (t) and the yield of the third overall flash for a delay time of 5 min, divided by the steady state oxygen flash yield.




RESULTS

Site-directed Mutagenesis of psbA Codon 170

The D1 polypeptide of PSII is encoded by the chloroplast gene, psbA, which in C. reinhardtii is a split gene containing five exons (Erickson et al.(1984); see Fig. 2). In vitro site-directed mutagenesis of pXb1.8, which contains psbA codon 170, was performed and four different mutant pXb1.8 plasmids were identified and characterized. The nucleotide substitutions introduced at psbA codon 170, the amino acid changes in the predicted protein product (aspartate histidine, asparagine, threonine, or proline), and restriction endonuclease sites introduced by the mutations are summarized in Fig. 2. The DNA sequence of the psbA exon 2 and exon 3 regions of each mutant pXb1.8 was determined (data not shown). Our results confirmed the presence of the mutation at codon 170 as shown in Fig. 2and verified that no other changes had been introduced inadvertently into the psbA coding regions contained in the mutant pXb1.8 constructs.

To facilitate chloroplast transformation of two different algal hosts, the mutant XbaI fragment from each mutant pXb1.8 was transferred into the larger plasmid constructs, pRR and pRRX (see Fig. 2). The resulting mutant pRR constructs (pRR-D170H, pRR-D170N, pRR-D170T, pRR-D170P) were used in co-transformation experiments in which the wild-type photosynthetic C. reinhardtii strains 2137 or 137y served as algal hosts. The mutant pRRX constructs (pRRX-D170H, pRRX-D170N, pRRX-D170T, pRRX-D170P) were used for all co-transformation experiments in which FuD7 was the algal host strain.

Chloroplast Transformation and Identification of Homoplasmic C. reinhardtii Co-transformants

Spectinomycin-resistant colonies arising on selective medium after biolistic bombardment of algal host cells with the spectinomycin resistance marker (pCrBH4.8) and a mutant psbA construct were picked and screened by colony hybridization where appropriate (see below) or by restriction fragment length polymorphism (RFLP) analysis. Eight independent spectinomycin-resistant colonies arising after co-transformation of wild-type hosts with pRR-D170T were tested, and two of these were heteroplasmic for the 240-bp BsrI fragment associated with the D170T mutation (Fig. 3A). All secondary colony isolates derived from these heteroplasmic transformant lines were homoplasmic with respect to this RFLP, as shown in Fig. 3B. Extended exposure of the blots revealed no evidence for mixed populations of fragments in any of the secondary isolates.

When the psbA deletion mutant FuD7 was used as the algal host for co-transformation, spectinomycin-resistant transformant colonies were screened by colony hybridization. Subsequent Southern blot analysis of DNA isolated from secondary isolates of original colonies which showed positive hybridization signals (data not shown) confirmed the presence of two copies of psbA per chloroplast genome. We were unable to detect any FuD7 host DNA molecules in these transformants using a polymerase chain reaction DNA amplification assay with the sensitivity to detect one copy of the FuD7 chloroplast DNA molecule in 2000, (data not shown) providing further evidence that the mutant lines are homoplasmic.

Table 1summarizes some of the mutant and control algal transformant lines isolated, the algal strains which served as host, and the transforming DNAs used. Three of the mutant transformants (D170N, D170T, D170P) are obligate heterotrophs. The D170H transformant grows very slowly on minimal medium.

To verify that the algal co-transformant lines contained the desired site-directed mutations, total RNA was isolated from transformant algal cells and used as a template for avian myeloblastosis virus reverse transcriptase in dideoxy DNA sequence reactions. All algal transformants showed the presence of the predicted mutations at codon 170 (Fig. 4), and no other differences were observed between the mutant and wild-type RNA templates. Finally, the mutant phenotype (poor or no growth on minimal medium) could be rescued by transformation of mutants with wild-type pXb1.8 DNA.


Figure 4: DNA sequence analysis using RNA templates isolated from mutant Chlamydomonas transformants. Total RNA isolated from wild-type (WT) and transformant cell lines served as a template for dideoxy sequence analysis using synthetic oligonucleotide primers complementary to psbA RNA as described under ``Experimental Procedures.'' An autoradiogram of an 8% acrylamide gel shows the region corresponding to DNA fragments 53-62 nucleotides in length. The sequence generated at wild-type (WT) codon 170 is shown to the left of the gel next to the complimentary sequence for the aspartate (Asp-170) GAC codon. Listed below the WT and each of the transformant sequences is the amino acid residue encoded at position 170, the codon sequence (5` to 3`), and the actual DNA sequence (3` to 5`) seen on the autoradiogram, respectively. AGCT above the WT sequence denotes the dideoxy sequencing reactions loaded in each of four consecutive wells, respectively.



In Situ Measurements of Fluorescence Emission

A simple and rapid, nonintrusive way to characterize PSII is to monitor the kinetics of chlorophyll fluorescence emission induced after illumination of C. reinhardtii cells on an agar plate with the lid in place (Bennoun and Delepelaire, 1982). In wild-type cells, the characteristic initial rise in fluorescence emission is due primarily to an accumulation of the reduced primary quinone acceptor Q(A) of PSII; the kinetics of Q(A) accumulation is dependent upon the relative rates of light-driven reduction of Q(A) and oxidation of Q(A) (for reviews, see Krause and Weis(1991), Horton and Bowyer(1992)). Fig. 5shows the variable component of the in situ fluorescence induction transients measured on algal host cells and the Asp-170 transformant cell lines. Each cell line in which codon 170 is altered exhibits a characteristic reduction in variable fluorescence. All the site-directed mutants show some variable fluorescence, in the relative amounts of WT > D170H > D170N > D170T > D170P. The host FuD7 exhibits none, as expected of a strain totally lacking PSII (Bennoun et al., 1986). Wild-type 2137 host cells or host FuD7 cells restored with wild-type psbA (FWT) showed kinetic traces similar to that of the 137y host. The presence of the spectinomycin resistance marker in the wild-type host (WTsr) or the FuD7 host restored with wild-type psbA (FWTsr) did not affect the fluorescence induction kinetics of these control strains (data not shown).


Figure 5: Kinetics of chlorophyll fluorescence emission from Chlamydomonas cells. Chlorophyll fluorescence induction transients were measured in situ from C. reinhardtii cells on algal plates dark-adapted for 10 min prior to assay, as described under ``Experimental Procedures.'' The relative amounts of PSII variable fluorescence in the FuD7 algal host and in isolates D170H(a), D170N(b), D170T(c), and D170P(b) of transformant cell lines are shown here. Initial levels of fluorescence were normalized to 0.2 for each strain. In the absence of normalization, the FuD7 host strain (which lacks PSII) displays a high level of initial fluorescence, approximately 5 times the absolute initial fluorescence of wild-type cells. Panel A, full time scale is 1.43 s; panel B, enlargement of the first 0.68 s of induction of selected algal strains as marked.



PSII Oxygen Evolution

The rate of steady state oxygen evolution by liquid cultures of C. reinhardtii transformants was measured as described under ``Experimental Procedures.'' Results of these measurements are reported in Table 1. The D170H and D170N transformants evolved oxygen at 30-60% and 10-20% of wild-type rates, respectively, while no oxygen evolution was detected in the D170T and D170P transformants. Rates of oxygen evolution were not altered by the presence of the spectinomycin resistance marker in control transformants of the wild-type host (WTsr) or control FuD7 cells restored with the wild-type psbA gene (FWTsr).

Quantitation of PSII Centers

To determine whether the reduction in (D170H, D170N) or loss of (D170T, D170P) PSII oxygen evolution measured in a population of cells in vivo was correlated with a commensurate loss of PSII, the amount of PSII present in transformants was estimated using a diuron binding assay (Vermaas et al., 1990). Several types of herbicides, including diuron (DCMU) and atrazine, bind at the Q(B) site of the D1 polypeptide of PSII with high affinity and in competition with the terminal quinone acceptor of PSII (Tischer and Strotmann, 1977; reviewed in Trebst(1991) and Oettmeier(1992)). There is one such high affinity binding site per PSII monomer, and the use of an assay which measures [^14C]diuron binding allows for quantification of the number of PSII centers, on a per chlorophyll basis, in whole cells (Vermaas et al., 1990). The assay also allows determination of the dissociation constant (K(D)) of the herbicide at its binding site. Results of PSII quantitation assays are shown in Fig. 6and summarized in Table 1. All of the mutants except D170P have the same levels of PSII centers as control cells. In the D170P mutant, PSII levels are reduced by approximately 50%. The K(D) for diuron is not significantly altered in any of the mutant transformants compared to wild-type and control transformant cells (Table 1), and the values observed are close to the 23 nM reported for Chlamydomonas thylakoids (Boschetti et al., 1985). These results suggest that the altered ability of the transformants to evolve oxygen is not caused primarily by a reduction in the number of PSII centers, but rather by a specific defect in donor-side function in the PSII centers which are present.


Figure 6: Quantification of PSII in mutant Chlamydomonas transformants. A [^14C]diuron herbicide binding assay (Vermaas et al., 1990) was used to quantitate PSII in whole cells. Assay of the number of diuron binding sites, assessed on a per chlorophyll basis, provides an accurate quantification of assembled PSII reaction centers. Assays were performed in duplicate, such that each point represents the mean of a pair of measurements for bound [^14C]diuron plotted against the mean of a pair of measurements for free [^14C]diuron, obtained after cells were incubated with labeled diuron at seven initial concentrations (ranging from 5-100 nM). The double reciprocal plot of 1/bound (mg of chlorophyll per nmol of [^14C]diuron bound) versus 1/free (1/µM [^14C]diuron free) for the nonaveraged data was used for extrapolation of chlorophylls/binding site at the y-intercept and -1/K at the x-intercept. The results are summarized in Table 1.



Evidence That Transformants Lack a Tertiary Electron Donor to P

A more detailed examination of PSII electron transfer was carried out using an assay in which algal cells in suspension are exposed to a series of short (2 µs) saturating flashes given 600 ms apart, and the kinetics of relaxation of the fluorescence yield of chlorophyll are measured after each flash (Nixon and Diner, 1992; Nixon et al., 1992b). Fig. 7shows traces obtained when this assay was applied to the C. reinhardtii transformants. The most striking results are obtained with the D170T transformant (Fig. 7E), in which the initial fluorescence yield measured 50 µs after the second actinic flash is reduced dramatically compared to that measured after the first flash, and initial fluorescence levels continue to decline with subsequent flashes. This is in marked contrast to control cells (Fig. 7A) which exhibit a high initial fluorescence yield at each flash. The initial fluorescence yield in D170P also declined significantly at the second, third, and subsequent flashes (Fig. 7D), while that of the D170H and D170N transformants declined more gradually with each successive flash (Fig. 7, B and C).


Figure 7: Relaxation of the chlorophyll fluorescence yield in mutant Chlamydomonas cells following saturating light flashes. The kinetics of relaxation of the chlorophyll fluorescence yield in whole cells was monitored following each of a series of saturating light flashes given 600 ms apart. C. reinhardtii control cells (panel A) and transformants (panels B-E) were suspended in TAP growth medium at a cell concentration close to 0.5 A and dark-adapted for 10 min. The amplitudes of the relaxation curves were normalized to what they would be if the cell concentrations were exactly 0.5 A for all cell suspensions. The earliest time point for each trace is at 50 µs following the actinic flash. A, FWTsr(a); B, D170H(a); C, D170N(a); D, D170P(b); E, D170T(a). The relative, normalized variable fluorescence (F - F(0)/F) is given on the y axis. Control and transformant strains are as described in the text and under ``Experimental Procedures.''



The most likely interpretation of the data is that, in dark-adapted D170T cells, for example, only a single electron is available on the donor side for reduction of P. It is assumed this electron comes from the tyrosine residue 161 (Y) of D1, resulting in formation of the radical species Y. In wild-type cells, P is reduced rapidly by an electron from Y which is rereduced in turn by the OEC, between actinic flashes. In such cells, the high fluorescence state is repeatedly observed 50 µs after each flash. Q is still dissipated by electron transfer to Q or by charge recombination with the donor side over longer time scales, resulting in the characteristic relaxation of the fluorescence yield of chlorophyll. However, in the D170T and D170P mutants, the oxygen-evolving complex which normally provides electrons for the reduction of Y is apparently absent or unable to function, leading to formation of YPPheoQ after the second flash. P is known to quench chlorophyll fluorescence (Butler et al., 1973) despite the presence of Q. Hence, the extent to which fluorescence quenching appears with each successive actinic flash is a reflection of the inactivity of the OEC. The greater the reduction in ability of a mutant to evolve oxygen, the more marked is the fluorescence quenching, again consistent with a functional defect in the tertiary electron donor (the OEC).

The rate of charge stabilization on the donor side of PSII in transformants can be inferred from experiments that monitor the rate of charge recombination between Q(A) and the donor side. In this assay, algal cells in suspension are treated with the inhibitor DCMU to block oxidation of Q(A) by Q(B) (see Fig. 1). After a single saturating light flash in the presence of DCMU, the rate of relaxation of the fluorescence yield of chlorophyll is, to a first approximation (ignoring energy transfer between PSII centers), the rate of reoxidation of Q(A) by charge recombination with the donor side. The rate of charge recombination depends on the concentrations of Q(A) and P; the higher the concentration of P, the faster the charge recombination (Nixon and Diner, 1992). Fig. 8shows the results of such studies. In the ``wild-type'' control line (FWTsr(a)), where the OEC is intact and the concentration of P is determined by the equilibrium S(1)Y(Z)P S(2)Y(Z)P (refer to Fig. 1legend), the fluorescence yield decays relatively slowly in a biphasic manner. Charge recombination in the D170H transformant is very similar to that seen in control cells. However, in the algal transformants D170N, D170P, and D170T, the rate constant for decay of the fast phase is approximately 3 to 4 times that observed for control cells. These results, consistent with our observations that the D170N, D170T, and D170P transformants are drastically reduced in their ability to evolve oxygen (Table 1) and exhibit a fluorescence quenching at 50 µs indicative of an increased concentration of P after two consecutive actinic flashes (Fig. 7), further strengthen our conclusion that the D170N, D170T, and D170P mutants are defective in PSII donor-side function.


Figure 8: Charge recombination between the PSII donor and acceptor sides in mutant Chlamydomonas cells. The kinetics of relaxation of the chlorophyll fluorescence yield in whole cells in the presence of 40 µM DCMU was monitored following a single saturating flash given to control (FWTsr(a)) and transformant (D170H(a), D170N(a), D170P(b), D170T(a)) cell lines. The cells were suspended in the same media and at the same cell concentrations as in Fig. 7. All curves were normalized to the same initial value at the first time point.



Oxygen-evolving PSII Complexes in the D170H Transformant Are Relatively Unperturbed

Since the D170H transformants evolve oxygen at 30-60% of the wild-type rate but accumulate wild-type quantities of PSII reaction centers, it was important to determine whether only some of these reaction centers were normally active or whether all reaction centers were active but showed perturbed function of the OEC. Use of the Joliot type oxygen electrode, in which a single layer of dark-adapted cells is sedimented onto the electrode surface, allows for the observation of oscillations in oxygen evolution produced after each flash in a series of consecutive saturating flashes as the active oxygen-evolving complexes cycle through the S-states in a synchronized manner (Fig. 9A). In light-grown wild-type cells, the majority of PSII centers relax to the S(1) state after a short dark incubation and thus the peak of oxygen evolution occurs on the third flash as the S(3) state is advanced to the S(4) state, resulting in release of oxygen and return to the S(0) state (see Fig. 1legend). In D170H transformant cells, peak oxygen evolution also occurred on the third flash. An estimation of the half-life time of the S(2) state was made based on measurements obtained using the Joliot electrode as described under ``Experimental Procedures'' and Fig. 9B legend. The results of such studies (Fig. 9B) show that the S(2) half-life time in the D170H mutant (10 s) is roughly comparable to that of the control cells (14 s). Thus, the PSII centers of D170H which are active in oxygen evolution appear to function quite normally.


DISCUSSION

C. reinhardtii is an ideal organism for studies involving the genetic engineering of chloroplast-encoded proteins which function in photosynthesis. This unicellular eukaryote contains a single large chloroplast providing a relatively large target for biolistic transformation, is able to grow heterotrophically on acetate allowing for the maintenance of strains defective in photosynthetic function, and has a well-characterized genetic system. Since the first report of biolistic chloroplast transformation (Boynton et al., 1988), this technique has been used to address many questions related to chloroplast function (reviewed in Boynton and Gillham (1993)), including studies focused on PSII structure and function (Przibilla et al., 1991; Roffey et al., 1991, 1994; Heiss and Johanningmeier, 1992; Lers et al., 1992; Schrader and Johanningmeier, 1992; Monod et al., 1994; Takahashi et al., 1994). The work we report here explores the role of the D1 polypeptide in activating chloroplast PSII donor-side function.

Site-directed Mutations at psbA Codon 170 Result in the PSII Phenotype of Homoplasmic Transformant Cell Lines

The single chloroplast of a C. reinhardtii cell contains 80-100 copies of the circular chloroplast DNA molecule (Harris, 1989), and each DNA molecule contains two identical copies of the D1 gene, psbA, which is located in the inverted repeat (Erickson et al., 1984). Integration of the transforming DNA into one or more chloroplast DNA molecule(s) of the host cell via homologous recombination (Newman et al., 1990) followed by segregation of the newly formed recombinant chloroplast DNA molecules during colony formation results in co-transformed cell lines which contain both the selectable marker and the unselected mutant psbA genes. Approximately 25% of our initial spectinomycin-resistant colonies were co-transformants which contained mutant copies of psbA, and all secondary colony isolates were homoplasmic for psbA (Fig. 3), regardless of whether a psbA deletion mutant host or wild-type algal host was used. Although independent integration events could account for the presence of two mutant psbA copies per chloroplast DNA molecule in transformants, it is likely that gene conversion and/or inter/intramolecular recombination maintains homogeneity of gene copies within the chloroplast-inverted repeat (Palmer, 1983; Erickson et al., 1984; Blowers et al., 1989). Treatment of algal host cells with 5-fluoro-2`-deoxyuridine (FdUrd) prior to bombardment reduces the number of copies of the chloroplast DNA molecule and increases the number of transformants recovered (Newman et al., 1990; Kindle et al., 1991). However, FdUrd is known to be mutagenic (Wurtz et al., 1979). For this reason, we do not treat cells with FdUrd and still are able to recover drug resistance transformants at a frequency of 10 to 10.

Several lines of evidence led us to the conclusion that the differences observed in the PSII phenotype of the mutant transformant cell lines are due to the specific mutations introduced at psbA codon 170 and not to any other factors. First, extensive molecular characterization of each different psbA plasmid construct used for transformation ruled out the possibility that other mutation(s) had been introduced inadvertently into the mutagenized region (pXb1.8) of psbA. Second, DNA sequence analysis using RNA templates isolated from each algal transformant cell line confirmed the presence of the site-directed mutations at psbA codon 170 and revealed no differences in any other region of the transcript analyzed. Third, all mutant cell lines, which grew very poorly or not at all on minimal medium, could be rescued to wild-type photosynthetic phenotype by transformation with a wild-type pXb1.8 construct, indicating that any mutations responsible for the altered photosynthetic phenotype were localized to the region of D1 contained in pXb1.8 (see Fig. 2). Fourth, the phenotypes of independent isolates of transformants bearing the same psbA mutation were similar (Table 1).^2 Finally, the fact that the PSII phenotypes of the D170T(b) (FuD7 host) and D170T(c) (wild-type host) cell lines were indistinguishable with respect to oxygen evolution, herbicide binding, and PSII quantitation ( Fig. 6and Table 1) as well as fluorescence measurements (Erickson et al., 1992) suggests that the nuclear genome of the host strain makes no significant contribution to the differences seen in PSII function in these mutants and strongly supports our conclusion that the alteration observed in PSII phenotype of a mutant is due solely to the amino acid substitution in the D1 polypeptide. We also demonstrated that the presence of the spectinomycin resistance marker present in all co-transformants had no effect on our assays of PSII function. Although this result was expected since initial characterization of the C. reinhardtii spectinomycin-resistant mutant spr-u-1-6-2, which contained the same 16 S rDNA mutation we used as a selectable marker, showed the mutation did not significantly alter photosynthesis or chloroplast protein synthesis (Chua and Gillham, 1977; see also Boynton and Gillham(1993)), we constructed and analyzed the ``control'' transformant algal strains WTsr, FWTsr, and FWT (see ``Experimental Procedures'').

Algal Transformants Accumulate Normal Levels of PSII Reaction Centers, but Are Defective in PSII Donor-side Function

It is known that the D1 polypeptide is essential for assembly and/or stability of chloroplast PSII (Bennoun et al., 1986), as is the D2 polypeptide (Erickson et al., 1986). In cyanobacterial systems, introduction of a wide range of single amino acid substitutions into the D1 or D2 polypeptides has resulted in the depletion or complete loss of PSII centers (for reviews, see Debus (1992), Nixon et al. (1992a), and Vermaas and Pakrasi (1992)). It is thus apparent that even small alterations to the primary structure of one of these two core PSII polypeptides can lead to destabilization of PSII, limiting the usefulness of such mutants in structure-function studies. Of particular interest for our studies was the question of whether our site-directed algal mutants contained PSII. Since all the transformants reported in this study assemble substantial levels of mutant PSII centers, the mutant PSII phenotype is due not to a lack of PSII reaction centers, but rather to a specific functional defect in the PSII centers present in the mutants.

The dramatic reduction in variable fluorescence in the mutants (Fig. 5) is correlated with loss of donor-side function (Table 1); the greater the reduction in variable fluorescence emitted by an algal mutant, the greater the reduction in rates of oxygen evolution. In situ monitoring of algal colonies for the kinetics of chlorophyll fluorescence induction should provide a rapid and effective method, in future experiments, for identifying mutants with specific defects in donor-side function.

A Role for D1 Aspartate Residue 170 in Assembly of a Functional Chloroplast Oxygen-evolving Complex

The C. reinhardtii mutants with amino acid substitutions at D1 residue 170 are partially or totally inhibited in oxygen evolution. Two types of fluorescence relaxation measurements ( Fig. 7and Fig. 8) allowed us to further assess electron donation to the PSII reaction center. Mutants in which D1 aspartate 170 is replaced by threonine (D170T) or proline (D170P) evolve no oxygen and display a progressively increased quenching of fluorescence yield observed at 50 µs following each in a series of consecutive saturating flashes (Fig. 7, D and E), as well as an accelerated rate of charge-recombination between the acceptor and donor sides of PSII (Fig. 8). Such results provide strong evidence for a loss of function of the OEC which acts as the tertiary donor to PSII. The catalytic site of the OEC is a cluster of 4 manganese ions which is assembled in a multistep process (see Tamura and Cheniae(1987), reviewed in Debus(1992)). Because light is required for oxidation of the first Mn atom bound at a high affinity site, as well as for oxidation of a second and subsequent ligated Mn, the term photoactivation has been applied to the process of assembly of the manganese cluster (see MiyaoTokutomi and Inoue(1992) for a recent discussion of photoactivation). Our results imply an inability of mutants to assemble a functional OEC and suggest that they may be defective in OEC manganese binding. In the green algal mutant Scenedesmus LF1 (Metz et al., 1980, 1986; Diner et al., 1988) and in cyanobacterial mutants with a modified C terminus (Nixon et al., 1992b), all of which lack a functional OEC but which contain aspartate at residue 170 of D1, the fluorescence yield also drops off at each successive flash, but much more gradually. This is presumably because Mn bound at aspartate 170 of D1 can provide electrons for the reduction of Z. Analysis of PSII core complexes depleted for manganese and isolated from Synechocystis sp. PCC 6803 mutants with amino acid substitutions at position 170 of D1 revealed an inverse relationship between the K(m) for the oxidation of exogenous Mn and the ability of the mutants to assemble active manganese clusters, providing further evidence for a role of this residue in the binding of manganese necessary for assembly of the oxygen-evolving complex (Nixon and Diner, 1992; Diner and Nixon, 1992). The apparent K(m) of 1 µM measured by these authors for the manganese oxidation site in wild-type cells was similar to the value observed for the site of photooxidation of the first manganese assembled during activation of the PSII OEC in vitro (Blubaugh and Cheniae, 1990), leading to the suggestion that aspartate 170 of D1 binds the manganese necessary for the first stage in assembly of the tetranuclear cluster.

When histidine or asparagine are substituted at residue 170 in D1, the resulting algal mutants (D170H, D170N) are partially inhibited in oxygen evolution and the kinetics of fluorescence relaxation are rather more complex. We attribute this phenotype to a heterogeneous population of PSII centers in these mutants, some with a functioning OEC and others without. While absolute rates of oxygen evolution in the D170N mutant were too low to allow a detailed investigation of oxygen-evolving PSII centers, active centers in the D170H transformant, which evolves oxygen at rates 30-60% those of wild-type cells, were analyzed using a Joliot type oxygen electrode. Such analysis revealed little perturbation in function of the OEC, with the S-states cycling relatively normally (Fig. 9A). The S(2) state decayed only slightly faster than in control cells, as judged by measurements of oxygen flash yields (Fig. 9B) and by the rate of charge recombination in the presence of DCMU (Fig. 8). Moreover, the fact that neither S(2)Q(B) recombination ( Fig. 9B) nor S(2)Q(A) recombination rates (Fig. 8) are greatly perturbed in the D170H transformant compared to the control cells indicates that the D1 substitution in D170H has no significant effect on the equilibrium constant for the reaction Q(A)Q(B) Q(A)Q(B) and suggests that PSII acceptor-side function is apparently unaltered in this mutant. Similar results have been obtained using a cyanobacterial mutant containing a histidine substitution at D1 residue 170 (Nixon and Diner, 1992). We suggest that the replacement of aspartate by histidine and, possibly asparagine, may lower the probability of successful completion of the earliest stage in OEC assembly, without affecting the performance of centers in which assembly of the manganese cluster is actually completed.

It is not yet clear whether aspartate 170 of the D1 polypeptide provides a ligand to manganese atoms in the mature tetranuclear manganese cluster of the activated OEC, whether this residue provides a ligand to manganese only during initial assembly of the manganese cluster, or whether aspartate 170 never provides a direct ligand to manganese, but rather has a critical effect on the conformation of the manganese binding site. The fact that different amino acid substitutions at residue 170 allow for wild-type OEC function in cyanobacterial mutants (Nixon and Diner, 1992; Boerner et al., 1992) suggests that residue 170 may not provide a ligand to manganese in the mature OEC. Relative differences in the overall rates of oxygen evolution by mutants in vivo appear to reflect differences in the relative effectiveness of the amino acid residue at position 170 to provide an initial ligand to manganese. In our algal mutants, histidine was more effective than asparagine (Table 1); in cyanobacteria, glutamate is more effective than histidine (Nixon and Diner, 1992). X-ray and magnetic resonance spectroscopy suggests that oxygen and some nitrogen atoms provide ligands to the manganese cluster, although evidence for chloride has been reported also (for reviews, see Diner et al.(1991) and Debus(1992)). Analysis of the D170H mutants, using magnetic resonance techniques to assay possible increased nitrogen ligation, may provide a means of resolving the role of D1 residue 170 in the mature OEC. Nevertheless, our results to date show that substitutions at D1 residue 170 affect the ability of chloroplast PSII centers to evolve oxygen and clearly show that this residue is critical for assembly and/or stability of the manganese cluster of the chloroplast OEC.

What Happens to PSII Centers Lacking an Active OEC?

Following exposure to low intensity light in vitro, PSII membrane fragments which lack the manganese cluster of the OEC accumulate pigment cations, and suffer subsequent damage to the PSII reaction center (Blubaugh et al., 1991) in a process known as low-light photoinhibition. Quenching of variable fluorescence accompanies low-light photoinhibition and such quenching may persist even after dark incubation due to the stable nature of some quenching pigment cations and the inability of damaged reaction centers to rapidly reduce the oxidized primary donor P. In our flash train experiments (Fig. 7), the initial fluorescence levels reached at the first flash are lower for all the Asp-170 mutants than for the control cells and are particularly low for D170T and D170P. This depression in initial fluorescence yield may result from a slowing of P reduction, and/or from accumulation of long-lived quenching species following growth of the mutants in low light.

The 50% reduction in the level of PSII observed when aspartate 170 is altered to proline suggests that, as well as leading to a complete loss of oxygen evolution, the mutation also may perturb the steady state balance between assembly and turnover of the PSII reaction-center complex. The D1 polypeptide displays a rapid, light-dependent turnover (Mattoo et al., 1984; Ohad et al., 1984) which has been the subject of intense study in recent years. Current models suggest that D1 is periodically damaged, is consequently removed from the PSII complex, and is replaced with a newly synthesized D1 restoring activity to the complex (for reviews, see Mattoo et al. (1989), Guenther and Melis(1990), and Prásil et al.(1992)). As damage is thought to result largely from the very strong oxidants generated during normal function of the PSII complex, attempts to improve photosynthetic efficiency are currently focused on the D1 turnover process itself (Barber and Andersson, 1992). The use of site-directed mutagenesis to manipulate chloroplast PSII electron transfer will aid in dissecting the molecular events of in vivo PSII activation and photodamage and should provide important insights as to how plants modulate photosynthetic function as light levels change.


FOOTNOTES

*
This work was supported by National Science Foundation Grants DCB-9006550 and MCB-9316915 (to J. M. E.) and DCB-9017526 (to B. A. D.). This research was also supported by National Institutes of Health Grant S07-RR07009-26RR. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom reprint requests should be addressed.

(^1)
The abbreviations used are: OEE, oxygen evolution enhancer; OEC, oxygen-evolving complex; kb, kilobase(s); bp, base pair(s); RFLP, restriction fragment length polymorphism; DCMU, 3-(3,4-dichlorophenyl)-1,1-dimethylurea; eq, equivalents.


ACKNOWLEDGEMENTS

We are grateful to Wim Vermaas (University of Arizona, Tempe, AZ) for kindly providing us with the [^14C]diuron used in these studies and to Karen Kindle (Cornell University, Ithaca, NY) for sending us the pCrBH4.8 plasmid.


REFERENCES

  1. Andersson, B., and Åkerlund, H.-E. (1987) in The Light Reactions (Barber, J., ed) pp. 379-420, Elsevier Science Publishers B.V., Amsterdam
  2. Arnon, D. (1949) Plant Physiol. 24, 1-15
  3. Barber, J., and Andersson, B. (1992) Trends Biochem. Sci. 17, 61-66 [CrossRef][Medline] [Order article via Infotrieve]
  4. Bennoun, P., and Delepelaire, P. (1982) in Methods In Chloroplast Molecular Biology (Edelman, M., Hallick, R. B., and Chua, N.-H., eds) pp. 25-38, Elsevier Science Publishers B.V., Amsterdam
  5. Bennoun, P., Spierer-Herz, M., Erickson, J. M., Girard-Bascou, J., Pierre, Y., Delosme, M., and Rochaix, J.-D. (1986) Plant Mol. Biol. 6, 151-160
  6. Blowers, A. D., Bogorad, L., Shark, K. B., and Sanford, J. C. (1989) The Plant Cell 1, 123-132 [Abstract/Free Full Text]
  7. Blubaugh, D. J., and Cheniae, M. (1990) Biochemistry 29, 5109-5118 [Medline] [Order article via Infotrieve]
  8. Blubaugh, D. J., Atamian, M., Babcock, G. T., Golbeck, J. H., and Cheniae, G. M. (1991) Biochemistry 30, 7586-7597 [Medline] [Order article via Infotrieve]
  9. Bockholt, R., Masepohl, B., and Pistorius, E. K. (1991) FEBS Lett. 294, 59-63 [CrossRef][Medline] [Order article via Infotrieve]
  10. Boerner, R. J., Nguyen, A. P., Barry, B. A., and Debus, R. J. (1992) Biochemistry 31, 6660-6672 [Medline] [Order article via Infotrieve]
  11. Boschetti, A., Tellenbach, M., and Gerber, A. (1985) Biochim. Biophys. Acta 810, 12-19
  12. Boynton, J. E., and Gillham, N. W. (1993) Methods Enzymol. 217, 510-536 [Medline] [Order article via Infotrieve]
  13. Boynton, J. E., Gillham, N. W., Harris, E. H., Hosler, J. P., Johnson, A. M., Jones, A. R., Randolph-Anderson, B. L., Robertson, D., Klein, T. M., Shark, K. B., and Sanford, J. C. (1988) Science 240, 1534-1537 [Medline] [Order article via Infotrieve]
  14. Burnap, R. L., Shen, J.-R., Jursinic, P. A., Inoue, Y., and Sherman, L. A. (1992) Biochemistry 31, 7404-7410 [Medline] [Order article via Infotrieve]
  15. Butler, W. L., Visser, J. W. M., and Simons, H. L. (1973) Biochim Biophys Acta 292, 140-151 [Medline] [Order article via Infotrieve]
  16. Chua, N.-H., and Gillham, N. W. (1977) J. Cell Biol. 74, 441-452 [Abstract]
  17. Church, G. M., and Gilbert, W. (1984) Proc. Natl. Acad. Sci. U. S. A. 81, 1991-1995 [Abstract]
  18. Critchley, C. (1985) Biochim. Biophys. Acta 811, 33-46
  19. Debuchy, R., Purton, S., and Rochaix, J.-D. (1989) EMBO J. 8, 2803-2809 [Abstract]
  20. Debus, R. J. (1992) Biochim. Biophys. Acta 1102, 269-352 [Medline] [Order article via Infotrieve]
  21. Debus, R. J., Barry, B. A., Sithole, I., Babcock, G. T., and McIntosh, L. (1988) Biochemistry 27, 9071-9074 [Medline] [Order article via Infotrieve]
  22. De Rose, V. J., Yachandra, V. K., McDermott, A. E., Britt, R. D., Sauer, K., and Klein, M. P. (1991) Biochemistry 30, 1335-1341 [Medline] [Order article via Infotrieve]
  23. Diner, B. A., and Nixon, P. J. (1992) Biochim. Biophys. Acta 1101, 134-138
  24. Diner, B. A., Ries, D. F., Cohen, B. N., and Metz, J. G. (1988) J. Biol. Chem. 263, 8972-8980 [Abstract/Free Full Text]
  25. Diner, B. A., Nixon, P. J., and Farchaus, J. W. (1991) Curr. Opin. Struct. Biol. 1, 546-554
  26. Erickson, J. M., and Rochaix, J.-D. (1992) in The Photosystems: Structure, Function and Molecular Biology (Barber, J., ed) pp. 101-177, Elsevier Science Publishers B.V., Amsterdam
  27. Erickson, J. M., Rahire, M., and Rochaix, J.-D. (1984) EMBO J. 3, 2753-2762
  28. Erickson, J. M., Rahire, M., Malnöe, P., Girard-Bascou, J., Pierre, Y., Bennoun, P., and Rochaix, J.-D. (1986) EMBO J. 5, 1745-1754
  29. Erickson, J. M., Whitelegge, J. P., Koo, D., and Boyd, K. (1992) in Current Research in Photosynthesis (Murata, N., ed) Vol. III, pp. 421-424, Kluwer Academic Publishers, Dordrecht
  30. Fernandez, E., Schnell, R., Ranum, L. P., Hussey, S. C., Silflow, C. D., and Lefebvre, P. A. (1989) Proc. Natl. Acad. Sci. U. S. A. 86, 6449-6453 [Abstract]
  31. Fichot, O., and Girard, M. (1990) Nucleic Acids Res. 18, 6162 [Medline] [Order article via Infotrieve]
  32. Forbush, B., Kok, B., and McGloin, M. P. (1971) Photochem. Photobiol. 14, 307-321
  33. Ghanotakis, D. F., and Yocum, C. F. (1990) Annu. Rev. Plant Physiol. Plant Mol. Biol. 41, 255-276 [CrossRef]
  34. Guenther, J. E., and Melis, A. (1990) Photosynth. Res. 23, 105-109
  35. Grunstein, M., and Hogness, D. S. (1975) Proc. Natl. Acad. Sci. U. S. A. 72, 3961-3965 [Abstract]
  36. Harris, E. H. (1989) The Chlamydomonas Sourcebook , Academic Press, Oxford
  37. Harris, E. H., Burkhart, B. D., Gillham, N. W., and Boynton, J. E. (1989) Genetics 123, 281-292 [Abstract/Free Full Text]
  38. Heiss, S., and Johanningmeier, U. (1992) Photosynth. Res. 34, 311-317
  39. Horton, P., and Bowyer, J. R. (1992) in Methods in Plant Biochemistry (Harbourne, J. B., and Dey, P., eds) Vol. IV, pp. 259-296, Academic Press, Oxford
  40. Ikeuchi, M. (1992) Bot. Mag. Tokyo 105, 327-373
  41. Johanningmeier, U., Bodner U., and Wildner, G. F. (1987) FEBS Lett. 211, 221-224 [CrossRef]
  42. Joliot, P., and Kok, B. (1975) in Bioenergetics of Photosynthesis (Govindjee, ed) pp. 387-412, Academic Press, New York
  43. Kindle, K. L., Richards, K. L., and Stern, D. B. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 1721-1725 [Abstract]
  44. Kok, B., Forbush, B., and McGloin, M. (1970) Photochem. Photobiol. 11, 457-475 [Medline] [Order article via Infotrieve]
  45. Krause, G. H., and Weis, E. (1991) Annu. Rev. Plant Physiol. Plant Mol. Biol. 42, 313-349 [CrossRef]
  46. Kunkel, T. A., Roberts, J. D., and Zakour, R. A. (1987) Methods Enzymol. 154, 367-382 [Medline] [Order article via Infotrieve]
  47. Lers, A., Heifetz, P. B., Boynton, J. E., Gillham, N. W., and Osmond, C. B. (1992) J. Biol. Chem. 267, 17494-17497 [Abstract/Free Full Text]
  48. Marder, J. B., Chapman, D. J., Telfer, A., Nixon, P. J., and Barber, J. (1987) Plant Mol. Biol. 9, 325-333
  49. Mattoo, A. K., Hoffman-Falk, H., Marder, J. B., and Edelman, M. (1984) Proc. Natl. Acad. Sci. U. S. A. 81, 1380-1385 [Abstract]
  50. Mattoo, A. K., Marder, J. B., and Edelman, M. (1989) Cell 56, 241-246 [Medline] [Order article via Infotrieve]
  51. Mayes, S. R., Cook, K. M., Self, S. J., Zhang, Z., and Barber, J. (1991) Biochim. Biophys. Acta 1060, 1-12
  52. Mayfield, S. P., and Kindle, K. L. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 2087-2091 [Abstract]
  53. Mayfield, S. P., Bennoun, P., and Rochaix, J.-D. (1987a) EMBO J. 6, 313-318 [Abstract]
  54. Mayfield, S. P., Rahire, M., Frank, G., Zuber, H., and Rochaix, J.-D. (1987b) Proc. Natl. Acad. Sci. U. S. A. 84, 749-753 [Abstract]
  55. Messing, J. (1983) Methods Enzymol. 101, 20-78 [Medline] [Order article via Infotrieve]
  56. Metz, J. G., Wong, J., and Bishop, N. I. (1980) FEBS Lett. 114, 61-66 [CrossRef]
  57. Metz, J. G., Pakrasi, H. B., Seibert, M., and Arntzen, C. J. (1986) FEBS Lett. 205, 269-274 [CrossRef]
  58. Metz, J. G., Nixon, P. J., Rögner, M., Brudvig, G. W., and Diner, B. A. (1989) Biochemistry 28, 6960-6969 [Medline] [Order article via Infotrieve]
  59. Miyao-Tokutomi, M., and Inoue, Y. (1992) Biochemistry 31, 526-532 [Medline] [Order article via Infotrieve]
  60. Monod, C., Takahashi, Y., Goldschmidt-Clermont, M., and Rochaix, J.-D. (1994) EMBO J. 13, 2747-2754 [Abstract]
  61. Nanba, O., and Satoh, K. (1987) Proc. Natl. Acad. Sci. U. S. A. 84, 109-112 [Abstract]
  62. Newman, S. M., Boynton, J. E., Gillham, N. W., Randolph-Anderson, B. L., Johnson, A. M., and Harris, E. H. (1990) Genetics 126, 875-888 [Abstract/Free Full Text]
  63. Nixon, P. J., and Diner, B. A. (1992) Biochemistry 31, 942-948 [Medline] [Order article via Infotrieve]
  64. Nixon, P. J., Chisholm, D., and Diner, B. A. (1992a) in Plant Protein Engineering (Shewry, P., and Gutteridge, S., eds) pp. 93-141, University Press, Cambridge
  65. Nixon, P. J., Trost, J. T., and Diner, B. A. (1992b) Biochemistry 31, 10859-10871 [Medline] [Order article via Infotrieve]
  66. Oettmeier, W. (1992) in The Photosystems: Structure, Function and Molecular Biology (Barber, J., ed) pp. 349-408, Elsevier Science Publishers B.V., Amsterdam
  67. Ohad, I., Kyle, D. J., and Arntzen, C. J. (1984) J. Cell Biol. 99, 481-485 [Abstract]
  68. Palmer, J. D. (1983) Nature 301, 92-93
  69. Philbrick, J. B., Diner, B. A., and Zilinskas, B. A. (1991) J. Biol. Chem. 266, 13370-13376 [Abstract/Free Full Text]
  70. Pr á s il, O., Adir, N., and Ohad, I. (1992) in The Photosystems: Structure, Function and Molecular Biology (Barber, J., ed) pp. 220-250, Elsevier Science Publishers B.V., Amsterdam
  71. Przibilla, E., Heiss, S., Johanningmeier, U., and Trebst, A., (1991) The Plant Cell 3, 169-174 [Abstract/Free Full Text]
  72. Rochaix, J.-D. (1978) J. Mol. Biol. 126, 597-617 [Medline] [Order article via Infotrieve]
  73. Rochaix, J.-D. (1992) Curr. Opin. Genet. Dev. 2, 785-791 [Medline] [Order article via Infotrieve]
  74. Rochaix, J. D., and Erickson, J. M. (1988) Trends Biochem. Sci. 13, 56-59 [CrossRef][Medline] [Order article via Infotrieve]
  75. Rochaix, J.-D., Mayfield, S., Goldschmidt-Clermont, M., and Erickson, J. (1988) in Plant Molecular Biology, A Practical Approach (Shaw, C. H., ed) pp. 253-275, IRL Press, Oxford
  76. Roffey, R. A., Golbeck, J. H., Hille, C. R., and Sayre, R. T. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 9122-9126 [Abstract]
  77. Roffey, R. A., Kramer, D. M., Govindjee, and Sayre, R. T. (1994) Biochim. Biophys. Acta 1185, 257-270 [Medline] [Order article via Infotrieve]
  78. Ruffle, S. V., Donnelly, D., Blundell, T. L., and Nugent, J. H. A. (1991) Photosynth. Res. 34, 287-300
  79. Rutherford, A. W., Zimmerman, J.-L., and Boussac A. (1992) in The Photosystems: Structure, Function and Molecular Biology (Barber, J., ed) pp. 179-229, Elsevier Science Publishers B.V., Amsterdam
  80. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd Ed, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York
  81. Sanford, J. C., Smith, F. D., and Russel, J. A. (1993) Methods Enzymol. 217, 483-509 [Medline] [Order article via Infotrieve]
  82. Schrader, S., and Johanningmeier, U. (1992) Plant Mol. Biol. 10, 251-256
  83. Schreiber, U. (1986) Photosynth. Res. 9, 261-272
  84. Shen, J. R., Ikeuchi, M., and Inoue, Y. (1992) FEBS Lett. 301, 145-149 [CrossRef][Medline] [Order article via Infotrieve]
  85. Stewart, A. C., Ljungberg, U., Åkerlund, H.-E., and Andersson, B. (1985) Biochim. Biophys. Acta 808, 353-362
  86. Svensson, B., Vass, I., Cedergren, E., and Styring, S. (1990) EMBO J. 9, 2051-2059 [Abstract]
  87. Svensson, B., Vass, I., and Styring, S. (1991) Z. Naturforsch. Sect. C Biosci. 46, 765-776
  88. Takahashi, Y., Matsumoto, H., Goldschmidt-Clermont, M., and Rochaix, J.-D. (1994) Plant Mol. Biol. 5, 779-788
  89. Tamura, N., and Cheniae, G. (1987) Biochim. Biophys. Acta 890, 179-194
  90. Tang, X.-S., Fushima, K., and Satoh, K. (1990) FEBS Lett. 273, 257-260 [CrossRef][Medline] [Order article via Infotrieve]
  91. Tang, X.-S., Diner, B. A., Larsen, B. S., Gilchrist, M. L., Jr., Lorigan, G. A., and Britt, R. D. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 704-708 [Abstract]
  92. Tischer, W., and Strotmann, H. (1977) Biochim. Biophys. Acta 460, 113-125 [Medline] [Order article via Infotrieve]
  93. Trebst, A. (1987) Z. Naturforsch. Sect. C Biosci. 42, 742-750
  94. Trebst, A. (1991) In Caseley, J. C., Cussans, G. W., and Atkin, R. K. (eds) in Herbicide Resistance in Weeds and Crops , pp. 145-164, ButterworthHeinemann, Oxford
  95. Vermaas, W. F. J., and Pakrasi, H. B. (1992) in The Photosystems: Structure, Function and Molecular Biology (Barber, J., ed) pp. 231-257, Elsevier Science Publishers B.V., Amsterdam
  96. Vermaas, W., Charité, J., and Shen, G. (1990) Biochemistry 29, 5325-5332 [Medline] [Order article via Infotrieve]
  97. Webber, A. N., Packman, L., Chapman, D. J., Barber, J., and Gray, J. (1989) FEBS Lett. 242, 259-262 [CrossRef]
  98. Wurtz, E. A., Sears, B. B., Rabert, D. K., Shepherd, H. S., Gillham, N. W., and Boynton, J. E. (1979) Mol. & Gen. Genet. 170, 235-242

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.