©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
A Subunit Interaction in Chloroplast ATP Synthase Determined by Genetic Complementation between Chloroplast and Bacterial ATP Synthase Genes (*)

(Received for publication, March 15, 1995; and in revised form, May 12, 1995)

Zugen Chen (1), Ashley Spies (1), Ray Hein (1), Xiaolan Zhou (1) (2), Brian C. Thomas (1) (2), Mark L. Richter (1), Peter Gegenheimer (1) (2) (3)(§)

From the  (1)Department of Biochemistry, the (2)Molecular Genetics Program, and the (3)Department of Botany, University of Kansas, Lawrence, Kansas 66045-2106

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES.
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

FF-ATP synthases utilize protein conformational changes induced by a transmembrane proton gradient to synthesize ATP. The allosteric cooperativity of these multisubunit enzymes presumably requires numerous protein-protein interactions within the enzyme complex. To correlate known in vitro changes in subunit structure with in vivo allosteric interactions, we introduced the subunit of spinach chloroplast coupling factor 1 ATP into a bacterial F ATP synthase. A cloned atpB gene, encoding the complete chloroplast subunit, complemented a chromosomal deletion of the cognate uncD gene in Escherichia coli and was incorporated into a functional hybrid F ATP synthase. The cysteine residue at position 63 in chloroplast is known to be located at the interface between and subunits and to be conformationally coupled, in vitro, to the nucleotide binding site >40 Å away. Enlarging the side chain of chloroplast coupling factor 1 residue 63 from Cys to Trp blocked ATP synthesis in vivo without significantly impairing ATPase activity or ADP binding in vitro. The in vivo coupling of nucleotide binding at catalytic sites to transmembrane proton movement may thus involve an interaction, via conformational changes, between the amino-terminal domains of the and subunits.


INTRODUCTION

The FF-type ATP synthases utilize the energy of a transmembrane proton gradient to drive the conversion of ADP plus P to ATP. The enzymes from chloroplasts, mitochondria, and bacteria exhibit a striking similarity in their overall structure, being composed of F (chloroplast CF),()a membrane-spanning proton channel complex, and F (chloroplast CF), a peripheral complex that contains the catalytic site(s) for ATP synthesis and hydrolysis. The F portion is composed of five subunits designated to in order of decreasing molecular weight and with a stoichiometry of (Nalin and Nelson, 1987). ATP synthesis in vivo is absolutely dependent upon continued translocation of protons through the F and F complexes. The isolated F complex, however, catalyzes only the reverse reaction of ATP hydrolysis. The minimal subunit assembly that efficiently supports ATP hydrolysis is the complex (Hu et al., 1993). The and polypeptides are arranged in an alternating array such that each subunit makes contact with two subunits to form a hexameric ring (Boekema and Bottcher, 1992; Abrahams et al., 1994). In addition, the subunits make direct contact with the subunit (Musier and Hammes, 1987) and with the (Engelbrecht et al., 1986) subunit. Six nucleotide binding sites are located at subunit interfaces. The putative catalytic sites are formed by residues with a minor contribution from subunit residues (Abrahams et al., 1994). The reverse is true for the remaining three sites, located predominantly on subunits and thought to play a regulatory role. CF is thus an allosteric enzyme whose catalytic function requires correct structural communication between subunits and at least the , , and subunits.

The amino acid sequences of both the and subunits, but especially of the subunits, are highly conserved among the different ATP synthases (Walker et al., 1985), strongly suggesting that they are structurally and functionally homologous. Indeed, we previously demonstrated (Richter et al., 1986) that the subunit isolated from spinach chloroplasts could be reconstituted into -deficient F from Rhodospirillum rubrum chromatophores to form a functional hybrid FF enzyme. When this evidence is considered along with other structural and enzymological studies (for review, see Penefsky and Cross(1991) and Boyer(1993)), little doubt remains that all FF-type ATP synthases share the same basic catalytic mechanism and essential allosteric intersubunit interactions.

Several important protein-protein interactions are suspected to involve the amino-terminal region of the polypeptide (approximately residues 20-95). For example, sensitivity to the fungal peptide tentoxin, an inhibitor of allosteric cooperativity in CF (Richter et al., 1986), is determined in part by the side chain of residue 83 (Avni et al., 1992). Recently, Colvert et al.(1992) determined that Cys is inaccessible in the assembled CF complex but that it is accessible to covalent modification in the isolated subunit. The simplest explanation was that Cys is buried in an interface. This conclusion was confirmed from the crystal structure of the mitochondrial F1 complex, which indicated that the residue that is equivalent to Cys of CF is located at an interface between adjacent amino-terminal -barrel domains of neighboring and subunits. Recent studies of Miki et al. (1994a, 1994b) have indicated that conformational processes involving these contacting domains are important for the coupling of transmembrane proton movement to ATP synthesis. Mills et al. (1995) have shown that binding of ADP, but not ATP, to the isolated subunit induces a significant change in the conformation of the amino-terminal domain. This change was detected as altered fluorescence of probes attached directly to Cys, greater than 40 Å from the ADP binding site. These observations suggest that allosteric interactions between the different nucleotide binding sites, as well as proton-driven changes in nucleotide binding affinities, may be mediated by the amino-terminal -barrel structures of adjacent and subunits. To further examine such interactions, we have examined the effects of mutations at residue 63 on nucleotide binding and catalytic properties of spinach chloroplast CF.

We previously described (Chen et al., 1992) a bacterial expression system in which large amounts of chloroplast subunit can be produced to facilitate site-directed mutagenesis investigations of assembly and conformational coupling among CF subunits. To address questions of subunit interaction among either homologous or heterologous F subunits in situ and to determine whether mutant polypeptides are functional, we have developed an in vivo complementation system in which a cloned spinach chloroplast atpB gene, encoding the CF subunit, complements an Escherichia coli chromosomal deletion of the cognate uncD gene. In this communication we describe the properties of the in vivo complementation system and its application to investigating mutations in codon 63 of the spinach chloroplast atpB gene.


EXPERIMENTAL PROCEDURES.

Materials

E. coli strain JP17, obtained from A. Senior (Lee et al., 1991), contains a deletion of the uncD gene corresponding to amino acids 20-332 of the F subunit. The genotype of JP17 is uncD, argH, entA, pyrE, recA::Tn10, tet. Plasmid pRPG31 (Gunsalus et al., 1982) contains an internal portion of the E. coliunc operon, including uncD ( subunit) and uncC ( subunit), inserted into pBR322 and transcribed from an endogenous plasmid promoter. Spinach chloroplast CF and subunit were prepared as described elsewhere (Hu et al., 1993; Richter et al., 1986). Antibiotics (ampicillin, tetracycline, and tentoxin) were purchased from Sigma. Tryptone and yeast extract were from Difco. All other chemicals were of the highest quality reagent grade available.

Computational

Putative ribosome binding sites were scored using results from the Perceptron neural network analysis of Stormo et al.(1982). The DNA sequence for 71 nucleotides surrounding each initiating AUG was represented as a 4 71 binary matrix, and the 71 4 weight matrix W71 was taken from Stormo et al.(1982). The scalar evaluation score is the inner product of the sequence matrix and the weight matrix. The software used is available from the authors. Oligonucleotide primers were designed with the aid of software from Scientific and Educational Software (State Line, PA); software from this source was also used for simulation, analysis, and graphic presentation of cloning manipulations. Sequence alignments were performed via the ``PhD'' profile alignment facility (Rost and Sander, 1993) and refined manually.

Mutagenesis

Oligonucleotide-directed mutagenesis was accomplished essentially as described before (Chen et al., 1992) using single-stranded DNA produced from plasmid pBS(-)XB3a in E. coli CJ236. Mutagenic primers were 20-23-mers with altered sequence (in lower case) at codon 63: -C63A, 5`-gcT; -C63V, 5`-gtT; -C63W, 5`-TGg. Mutations were confirmed by partial sequencing. For overexpression, an XbaI-SstI fragment containing each mutation was used to replace the corresponding region of expression plasmid pET3a-NE3 (Chen et al., 1992).

Plasmid Construction

Plasmids pBS(-)XE0 and pET3a-NE2 have been described previously (Chen et al., 1992). pBS(-)XE0 contains an XbaI-EcoRI fragment of spinach chloroplast DNA bearing the atpB gene ( subunit) and the proximal 50 nucleotides of the overlapping atpE gene ( subunit). pET3a-NE2 contains atpB fused to the gene 10 ribosome binding site and translational enhancer of coliphage T7 (Olins and Rangwala, 1990) in the expression vector pET3a (Studier et al., 1990). The atpB insert and T7 ribosome binding site of pET3a-NE2 were excised with XbaI and BamHI and inserted into similarly cleaved pBS(-) (Stratagene) to produce plasmid pBS(-)XB1. The plasmid was digested with HindIII for 4 h at 37 °C in 50 mM KOAc buffer (1-4-All Plus buffer, Pharmacia Biotech Inc.) and filled in by the addition of E. coli DNA polymerase Klenow fragment (25 units/ml plus 0.5 mM dNTP for 50 min at 20 °C). The reaction mixture was extracted with phenol/chloroform, ethanol-precipitated, and self-ligated with T4 DNA ligase (Life Technologies, Inc.) overnight at 18 °C. DNA was transformed according to Hanahan(1985) into E. coli DH5 (Life Technologies Inc.) and was selected on LB medium (1% (w/v) NaCl, 1% (w/v) Tryptone, 0.5% yeast extract, pH 7.5) plus 100 µg of ampicillin/ml. Two (of about 50) of the recovered colonies were chosen and designated pBS(-)XB3a and -b. As documented below, both of these plasmids could express a functional subunit in E. coli. All subsequent experiments and constructions reported here as XB3 were performed with XB3a.

Complementation Assay

Plasmids were transformed into E. coli JP17 and plated onto LB agar plus 100 µg of ampicillin/ml and 12.5 µg of tetracycline/ml. Vector pBS(-) was also transformed into E. coli DH5 and selected without tetracycline. After growth overnight at 37 °C, colonies were streaked onto a prewarmed minimal medium plate containing succinate (Hawthorne and Brusilow, 1986; Gibson et al., 1977) (100 mM potassium phosphate (pH 7.1), 1 mM MgCl, 0.2 µM ZnSO, 0.2 µM CuSO, 0.5 µM MnSO, 2 nM CoCl, 0.5 µM FeSO, 10 µM CaCl, 0.1% (w/v) thiamine HCl, 40 µM 2,3-dihydroxybenzoate, 0.8 mM arginine, 0.2 mM uracil, 0.2% (NH)SO, 0.06% casein, 0.4% sodium succinate). For growth yield determinations, cells containing plasmids were inoculated into 3 ml of LB medium plus 100 µg of ampicillin/ml and (except for DH5) 12.5 µg of tetracycline/ml and grown overnight at 37 °C. A loop of each culture was transferred to 15 ml of prewarmed minimal medium containing succinate at 3, 6, or 12 mM plus appropriate antibiotics as above. Cell densities were measured as turbidity at 590 nm after growth for varying times at 37 °C.

ATPase Assays

Cells were grown on LB medium containing 30 mM glucose. Starter cultures (3 ml) were grown at 37 °C to an optical density at 590 nm of 0.7 then inoculated into 1 liter of the same medium and grown to midexponential phase. Cells were harvested by centrifugation for 15 min at 10,000 g, resuspended, and washed once in TM buffer (50 mM Tris-HCl (pH 8), 10 mM MgCl). The cells were finally suspended to 40% (wet w/v) in TM buffer and lysed at 4 °C with 0.1-mm glass beads in a Mini-Bead Beater (Biospec Products) at full speed. Unbroken cells and cell debris were removed by centrifugation at 10,000 g for 15 min. Plasma membranes were collected by centrifugation at 100,000 g for 3 h at 4 °C and resuspended to 10 mg of protein/ml. ATPase activity of washed membranes was determined by measuring release of P from ATP at 37 °C (Taussky and Shorr, 1953) in an assay mixture (1 ml) containing 50 mM Tris-HCl (pH 8), 4 mM ATP, 2 mM MgCl, and membranes equivalent to 50 µg of protein. The protein concentration of membrane preparations was measured by the BCA method (Pierce Chemical Co.) (Smith et al., 1985).

Immunoblot Analysis

From 10,000 g supernatants prepared as above, plasma membranes were obtained by centrifugation at 100,000 g for 1.5 h and washing once in 4 ml of TM buffer. The F complex was released by resuspending membranes in 7.5 ml of stripping buffer (1 mM Tris-HCl, 0.5 mM Na-EDTA, 2 mM dithiothreitol (pH 8)) and mixing gently overnight at 4 °C. The membranes were sedimented at 100,000 g for 1 h. Proteins were precipitated from the supernatant by the addition of trichloroacetic acid to 8% (Schumann et al., 1985). Proteins were solubilized in 1 ml of sample buffer (1.25% (w/v) sodium dodecyl sulfate, 7.5% sucrose, 125 mM dithiothreitol, 125 mM NaCO), and polypeptides were separated on 12% polyacrylamide gels (Schumann et al., 1985). One of a set of duplicate gels was stained with Coomassie Blue G-250; the other was electrotransferred to polyvinylidene difluoride membranes (Immobilon, Millipore) in a Hoefer TE apparatus following the manufacturer's instructions. The membrane was decorated overnight with a mouse monoclonal antibody (1:2500 dilution) raised against the purified chloroplast subunit. Decorated protein bands were reacted for 1 h with goat anti-mouse peroxidase-conjugated antibody (1:1000 dilution; Sigma).

Nucleotide Binding Assays

polypeptide was over-expressed in E. coli, purified, and refolded essentially as before (Chen et al., 1992). 0.5 mg of inclusion body protein, suspended at 0.1 mg/ml in TE50/2 buffer (50 mM Tris-HCl (pH 8), 2 mM Na-EDTA), was mixed with 150 ml of refolding buffer (20 mM Tris-HCl (pH 8), 2 mM Na-EDTA, 2 mM dithiothreitol) containing 4.3 M urea for 90 min at 4 °C, and then dialyzed successively against (i) 150 ml of refolding buffer plus 3 M urea, 12% glycerol for 5 h at 4°; (ii) 150 ml of refolding buffer plus 1.5 M urea, 10% glycerol for 7 h at 4°; (iii) 200 ml of refolding buffer plus 3 mM ATP, 8% glycerol for 6 h at 20°. Isolated polypeptides were titrated with trinitrophenyl-ADP (TNP-ADP). The net fluorescence enhancement was corrected for inner filter effect, and binding constants were determined as before (Chen et al., 1992; Mills and Richter, 1991) by least-squares nonlinear curve-fitting to the untransformed data. Since each subunit has one ADP binding site, the fraction of polypeptide which was correctly refolded was taken as the calculated fractional number of binding sites.


RESULTS

Sequence of the Spinach Chloroplast atpB Gene

An initial alignment of the spinach chloroplast amino acid sequence deduced from the gene sequence (Zurawski et al., 1982) (Genbank entry SPICPATBE, accession J01441) revealed several alterations in residues completely or highly conserved in all other FF-type polypeptides (see also Chen et al., 1992). To resolve these discrepancies, we resequenced most of the remainder of atpB. Three additional errors in the published sequence were located at positions G (was C), G (was C), and G (was A). These amendments change three amino acids: Ala (was Pro), Gly (was Arg), and Asp (was Asn). Each change results in greater sequence conservation with other polypeptides. The deduced amino acid sequence is presented in Fig. 1A, aligned with representative chloroplast, mitochondrial, and bacterial sequences. A more detailed comparison of the amino-terminal domain is given in Fig. 1B. The revised DNA sequence is GenBank accession U23082, locus SOU23082.


Figure 1: Deduced amino acid sequence of the spinach chloroplast CF subunit. A, the deduced amino acid sequence (GenBank accession U23082) is aligned over representative sequences of CF from a monocot gramineae (Oryzasativa), a bryophyte (Marchantiapolymorpha), a green alga (chlorophyte, oocystacea; Chlorellaellipsoidea, Swiss-Prot entry ATPB_CHLEL), and a cyanobacterium (Synechococcus PCC 6301), and of F ATP synthase from a purple non-sulfur bacterium (E. coli), and bovine mitochondrion. An initial alignment was performed with all chloroplast, mitochondrial, and bacterial F sequences in the SWISS-PROT data base; a representative subsampling is shown. Swiss-Prot entries not given are listed below. In sequences other than spinach, upper case letters denote nonconservative changes, whereas lower case letters indicate conservative or semiconservative changes (see Bordo and Argos(1991)). B, alignment of spinach residues 19-95 corresponding to the amino-terminal -barrel of bovine mitochondrial F (residues 9-80). The spinach chloroplast sequence is aligned with sequences from organisms representative of a uniformly broad range of taxa. The uppernumbers designate positions in the spinach chloroplast sequence, and the lowernumbers correspond to the bovine mitochondrial sequence. The unshadedareas represent the approximate extents of the six -strands identified in the crystal structure of bovine mitochondrial F (Abrahams et al., 1994). Sequences are identified by their Swiss-Prot entry names (given in parentheses below). Kingdom Eucarya: Tracheophytes - Angiosperms, Dicot, Solanaceae, tobacco Nicotianatabacum (ATPB_TOBAC) and Nicotianaplumbaginifolia (ATPB_NICPL); Dicot, Fabaceae, pea Pisumsativa (ATPB_PEA); Monocot, Gramineae, rice Oryzasativa (ATPB_ORYSA); Monocot, Poaceae, Aegilopscolumnaris (ATPB_AEGCO). Gymnosperm, Pinaceae, black pine Pinusthunbergii (extracted from Genbank accession PINCPTRPG). Pteridophyte, filicophyta, turnip fern Angiopterislygodiifolia (ATPB_ANGLY). Bryophyte: Hepaticopsida, liverwort Marchantiapolymorpha (ATPB_MARPO). Phycophytes: chlorophyte (green alga), chlamydomonadaceae, Chlamydomonasreinhardtii (ATPB_CHLRE); phaeophyte (brown alga) Pylaiellalittoralis (ATPB_PYLLI); ``prochlorophyte'' Prochlorondidemni (PIR entry A42697); ``euglenophyte'' Euglenagracilis Z (ATPB_EUGGR). Kingdom Bacteria: Cyanobacterium Synechococcus PCC 6301 (ATPB_SYNP6); Green sulfur bacterium Chlorobiumlimicola (ATPB_CHLLI); Purple sulfur bacterium Rhodospirillumrubrum; Purple non-sulfur bacterium E.coli (ATPB_ECOLI), plant mitochondrion (N. plumbaginifolia, ATP2_NICPL). (Mitochondria arose from the purple non-sulfur lineage of bacteria.)



Complementation of an E. coli uncD Deletion by the Spinach Chloroplast atpB Gene

To establish an in vivo system in which chloroplast could be produced in soluble form, we first examined the relevant sequences of spinach chloroplast atpB for potential E. coli promoters or ribosome binding sites. No likely matches to these sequences were found. In particular, atpB does not have a good E. coli Shine-Dalgarno sequence upstream. We applied the neural network predictor of Stormo et al.(1982) to estimate whether translational initiation would occur at the first AUG of atpB. Evaluation with weight matrix W71 (Stormo et al., 1982) predicted that atpB would be translatable in E. coli. Munn et al.(1991) had previously demonstrated that spinach chloroplast atpB was translated in E. coli minicells but that translation yields were increased by introducing a consensus E. coli Shine-Dalgarno sequence. We therefore tested an existing construct pBS(-)XB0 (Chen et al., 1992) in which atpB is preceded by native chloroplast DNA sequences. We also placed atpB under control of the E. colilacP promoter and provided it with a strong bacterial translation start signal; since we had also constructed a high-expression plasmid, pET3a-NE3, consisting of atpB fused to the bacteriophage T7 ribosome binding site and translational enhancer (Chen et al., 1992), we subcloned this T7-atpB fusion into plasmid pBS(-), giving plasmid pBS(-)XB1. Fig. 2shows the relevant features of these constructs. Plasmids XE0 and XB1 are high-copy-number derivatives of pUC19 and exhibit constitutive expression of the lac promoter as a result of titration of endogenous lac repressor. Cells containing these plasmids, particularly when grown in the absence of glucose, thus synthesize polypeptide constitutively (data not shown). In order to prevent read-through translation from lacZ into atpB, we modified plasmid pBS(-)XB1 by filling in a HindIII site at lacZ codon 18 to introduce in-frame termination 63 base pairs upstream of atpB. Upon further examination of the resultant plasmids pBS(-)XB3a and -b, however, we found that each contained, instead of the expected fill-in, a deletion extending from the HindIII site to one of two upstream HindIII* sites (AtGCTT and AgGCTT respectively) located within the lacP promoter. This region is depicted in Fig. 2B, and the sequences are shown in Fig. 2C. The deletion in plasmid XB3a starts just upstream of the -10 box of the lacP (P1) promoter (Reznikoff and McClure, 1986), whereas the deletion in XB3b also removes the -35 region of lacP. Transcription of atpB in these plasmids is evidently driven by constitutive promoters in the vector, such as the upstream P in the colE1 ori region (Balbas et al., 1988).


Figure 2: Expression clones of chloroplast atpB.A, plasmid pBS(-)XE0. Hatchedbars represent vector sequences; darkshading represents chloroplast atpB coding sequences; lightshading represents chloroplast atpE sequences; and openbars or arrows represent noncoding chloroplast insert sequences. Also indicated are the positions of HindIII and HindIII* restriction recognition sites, the pBS-derived lacP promoter, and the codon for Cys, discussed in the text. B, plasmids pBS(-)XB1 and pBS(-)XB3a and b. Shading is the same as in A, with the heavysolidline representing phage T7 sequences. The italicizedrestrictionenzymesites are derived from T7 or adaptor sequences. The figure shows the extent of the two deletions in plasmids XB3a and b. Plasmid pBS(-)XB1 is identical but lack the deletion; the leftmost vector region is intact up to XbaI, as in panelA. Position of the pET3a-derived T7 ribosome binding site (RBS) is indicated. C, sequence of the lac promoter region and deletions. Nucleotide and inferred amino acid sequences are shown for the lacP1 promoter, -galactosidase -peptide, and atpB translation initiation region from plasmids pBS(-)XB1 and the deletion derivatives pBS(-)XB3a and XB3b. Restriction site sticky ends are underlined; sequences created by filling-in are lowercase; and promoters and translation initiation codons are doubleunderlined.



Expression of atpB in cells transformed with these plasmids was monitored indirectly by an assay that requires formation of a functional ATP synthase in vivo. E. coli cells lacking ATP synthase are unable to respire and hence cannot use citric acid cycle intermediates such as succinate as an energy source. The genetic complementation assay thus measures the ability of an introduced gene to support bacterial growth on minimal medium containing succinate as the sole carbon source. The ATPase host is E. coli JP17 (Lee et al., 1991), which has a targeted deletion of most of the uncD gene encoding the F subunit. We transformed JP17 with plasmids either lacking an insert, containing the E. coli uncD gene, or containing the various constructs of spinach chloroplast atpB described above. After growth overnight on LB agar to allow expression of the inserted gene, cells were transferred to minimal agar containing 0.4% succinate. Complementation was scored as cell density after 1-3 days at 37 °C. As shown in Fig. 3, the uncD strain transformed with vector only (pBS(-)) fails to grow. As a positive control, cells containing E. coli uncD (plasmid pRPG31) (Gunsalus et al., 1982) grow almost as well as the wild-type strain DH5 transformed with vector. The chloroplast atpB gene in plasmid pBS(-)XB0, with constitutive transcription but native chloroplast translation control sequences, supports weak but detectable growth on solid media. Finally, Fig. 3shows that plasmid pBS(-)XB3, containing atpB with a weak plasmid promoter but a strong bacteriophage translational control region, supports growth almost as well as does the native E. coli gene. We were surprised to find, however, that the constitutively-transcribed, translationally-enhanced gene in plasmid pBS(-)XB1 could not complement the uncD deletion (not shown). Further investigation revealed that although this plasmid directs synthesis of reasonable amounts of polypeptide, the protein aggregates entirely into insoluble inclusion bodies (data not shown).


Figure 3: Complementation of an uncD deletion by cloned subunit genes. Colonies of JP17 (uncD) or DH5 cells transformed with the indicated plasmids were streaked onto succinate minimal plates as described under ``Experimental Procedures'' and grown for 3 days.



Assembly and Activity of Hybrid ATP Synthase

To provide a more quantitative assessment of ATP synthesis capability in bacteria carrying a chloroplast CF subunit gene, we determined the yield of cells grown on limiting concentrations of succinate, which is an indirect measure of ATP synthesis efficiency. To confirm that active ATP synthase had been assembled in these cells, we determined the specific ATPase activity of partially purified ATP synthase, and we identified chloroplast by Western blot analysis. The results shown in Table 1demonstrate that all strains capable of growth possess comparable levels (within one standard deviation) of Mg-dependent membrane-bound ATPase activity. Membranes from the uncD deletion mutant JP17 transformed with vector only exhibited undetectable ATPase activity, whereas cells transformed with E. coliuncD exhibited 86% of wild-type Mg-ATPase activity. Strikingly, chloroplast atpB restored Mg-ATPase activity to about 75% of the level achieved with the E. coliuncD gene or to about 65% of wild-type activity. Both the hybrid and the endogeneous enzymes exhibited only low levels of Ca-dependent ATPase activity (Table 1).



To confirm that this ATPase activity was that of a hybrid F species, washed membranes from selected transformants were treated with EDTA to release the F portion of ATP synthase. The presence of bacterial or chloroplast polypeptides in the released protein fraction was assayed by polyacrylamide gel electrophoresis and Western immunoblotting, as shown in Fig. 4. Strain JP17 transformed with vector only has no membrane-bound -reactive material (Fig. 4B, lane3), whereas when transformed with E. coli uncD it contains the 50.3-kDa E. coli (lane2). Membranes from JP17 transformed with clone XB3 (lane4) contain substantial amounts of the 53.6-kDa chloroplast polypeptide.


Figure 4: Wild-type and mutant subunit assembly into membrane-bound ATP synthase. Soluble protein released from washed membrane preparations was prepared, and polypeptides were fractionated by denaturing SDS-polyacrylamide gel electrophoresis, as under ``Experimental Procedures.'' Lane1, 20 µg CF; lanes2-6, 60 µg of membrane-eluted protein from E. coli JP17 transformed with the following plasmids: lane2, pRPG31; lane3, pBS(-), lane4, pBS(-)XB3; lane5, pBS(-)XB3-C63A; lane6, C63V; lane7, C63W. A, polypeptides were visualized by staining with Coomassie Blue. B, an identical gel was transferred to polyvinylidine difluoride membrane, and polypeptide was detected with an anti- monoclonal antibody. The upperband corresponds to spinach chloroplast (M = 53,571), and the lowerband corresponds to E. coli (M = 50,331).



Previous studies (Richter et al., 1986) showed that the CF subunit, when reconstituted in vitro with the -less F of R. rubrum chromatophores, conferred upon the hybrid enzyme sensitivity to the CF-specific inhibitor tentoxin. The hybrid enzyme also exhibited an enhanced dependence of Mg-ATPase activity on the oxyanion sulfite, compared with that of the R. rubrum F. We therefore examined the activity of the hybrid chloroplast-E. coli F in the presence of tentoxin or sulfite. Table 2demonstrates that neither the bacterial nor the hybrid membrane-bound ATPase preparation is inhibited by tentoxin concentrations well above those required to give almost total inhibition of native CF (see Hu et al.(1993)). Table 2also shows that low concentrations of sulfite, 2-10 mM, slightly stimulate (by 20-25%) the activity of the hybrid enzyme but not that of the bacterial enzyme. Higher concentrations of sulfite lead to significant inhibition of both enzymes.



Hybrid ATP Synthases with Amino Acid Replacements at CF Residue 63

The region surrounding Cys is of particular interest because it is exposed only upon removal of the polypeptide from the CF complex and is thus a candidate for a residue involved in intimate protein-protein interactions at the subunit interface (Colvert et al., 1992). The amino acid sequence from 60 to 70 (spinach numbering) is virtually identical among all chloroplasts and cyanobacteria and exhibits strong conservation with purple bacteria and mitochondria; these sequences are presented in Fig. 1B. To investigate the properties of this region, residue 63 in plasmid pET3a-XB3 was changed from Cys to Ala, Val, or Trp by oligonucleotide-directed mutagenesis to produce plasmids pET3a-XB3/C63A, C63V, and C63W.

To assess the functionality of these altered subunits in situ, the mutant genes were tested for their ability to complement the uncD deletion in E. coli strain JP17. Complementation was scored initially by a plate assay (as in Fig. 3) and then by measuring growth yields in succinate-limited medium. These in vivo data were corroborated with ATPase activities of isolated bacterial membranes. As seen in Table 3, the -less strain JP17 transformed with a plasmid encoding -C63A grows somewhat better than JP17 containing wild-type CF , and exhibits identical ATPase activity. JP17 containing -C63V grows less well, and JP17 containing -C63W exhibits essentially no growth after 32 h, a time by which JP17 transformed with wild-type is at stationary phase. (Some growth of JP17/-C63W was evident after 48 h.) In contrast to its poor growth, mutant -C63W shows near normal in vitro ATPase activity (90% versus wild-type). The native chloroplast DNA clone XE0, which supported growth on solid medium (Fig. 3), did not support growth in liquid medium (Table 3). A plot of growth yield versus [succinate], shown in Fig. 5A, reveals a linear dependence for JP17 transformed with every plasmid except the mutant -C63W, indicating that for -C63W a factor other than succinate is growth-limiting. We infer that in JP17/-C63W, ATP synthesis is growth-limiting. Furthermore, decreased growth yield is also directly correlated with increased bulk of the amino acid side chain of residue 63. As seen in Fig. 5B, a plot of growth yield versus side chain volume yields an inverse linear relationship.




Figure 5: Dependence of cell growth on [succinate] and side chain volume at residue 63. A, dependence of growth on [succinate]. Strain JP17 was transformed with clones of wild-type atpB (Cys) or the mutants C63A (Ala), C63V (Val), or C63W (Trp). Cell density was measured as turbidity at 590 nm after 32 h of growth. Each value is the mean of two determinations. B, dependence of cell growth on side chain volume of the amino acid at position 63. Growth yields for JP17 containing the four plasmids shown in panelA were determined at 32 h and 6 mM succinate. Absorbances were normalized to that of JP17/C63A. Amino acid side chain volumes were calculated directly from the crystallographic data compiled by Richards(1974); the volume of cysteine was taken as one-half cystine plus an average hydrogen volume of 7 Å. Very similar curves were obtained for different growth times and succinate concentrations.



That a true hybrid F had been assembled in these strains was confirmed by immunoblot analysis of F isolated from washed bacterial membranes. As seen in Fig. 4, lanes5-7, JP17 transformed with plasmids containing -C63A, -C63V, or -C63W contains substantial amounts of authentic membrane-bound spinach chloroplast polypeptide, comparable with the amount present in JP17 transformed with wild-type chloroplast (lane4) or with E. coli (lane2).

To determine whether the defect in ATP synthesis resulted from a defect in substrate binding, wild-type atpB and the mutant atpB genes C63A and C63W were expressed in E. coli BL21[DE3]/pLysS (Chen et al., 1992; Rosenberg et al., 1987; Studier et al., 1990), and the resultant polypeptides were refolded from purified inclusion bodies. Each subunit preparation was assayed for ADP binding by a standard titration with the fluorescent ADP analogue TNP-ADP. The dissociation constant, K, and binding site occupancy were calculated from duplicate or triplicate titrations. A typical result is shown in Fig. 6. As summarized in Table 4, both the C63W and C63A mutant subunits refolded to essentially the same extent as wild-type . The C63A subunit binds TNP-ADP with an affinity very similar to that of wild-type, whereas the C63W subunit binds TNP-ADP about twice as tightly. Since the C63W mutant subunit has close to normal nucleotide binding ability as well as ATP hydrolysis activity, the defect in ATP synthesis in vivo is unlikely to result from any intrasubunit abnormality.


Figure 6: Nucleotide binding to over-expressed wild-type and mutant CF . Binding titrations were performed and analyzed as under ``Experimental Procedures.'' One representative trial is shown. The corrected fluorescence enhancement was normalized to 180 µg (3.36 nmol) polypeptide.






DISCUSSION

Assembly In Vivo of a Functional Chloroplast-Bacterial Hybrid ATP Synthase

Although we had shown previously that high-level bacterial expression of the spinach chloroplast atpB gene leads to production of insoluble protein (Chen et al., 1992), we expected that moderate rates of expression might yield soluble protein in vivo. Furthermore, we had reason to hope that interspecies assembly of hybrid F ATP synthase would ensue, as functional ATP synthase has been produced in vitro by introducing chloroplast subunits into R. rubrum ATP synthase (Richter et al., 1986). Our results demonstrate that such a hybrid system can be formed and that it functions well enough to support vigorous bacterial growth. Surprisingly, every clone that appropriately expressed soluble chloroplast polypeptide to support cellular growth contained either an active promoter and a very weak ribosome binding site or a weak promoter and a strong ribosome binding site. In contrast, a clone in which atpB was transcribed from a strong (lacP) promoter and translated from a strong (phage T7) ribosome binding site expressed atpB only as an insoluble polypeptide. From such a construct, two spontaneous deletions of the lac promoter were isolated that allowed functional expression of atpB as soluble protein. Consistent results were obtained by Engelbrecht and co-workers (Lill et al., 1993; Burkovski et al., 1994), who found that spinach atpB expressed behind a phage promoter yielded entirely insoluble polypeptide and could not complement E. coliuncD or uncD mutations. These results all support the hypothesis that low-level expression from weak, endogenous plasmid promoters permits formation of correctly folded, functional CF polypeptide. Likewise, functional E. coli polypeptide is produced from plasmid pRPG31 by transcription of the E. coliuncD gene from a constitutive weak vector promoter, most likely the P1 promoter of tetR (Balbas et al., 1988).

The hybrid spinach/E. coli enzyme was sufficiently active in ATP synthesis to allow cells to grow on succinate almost as well as cells containing the native bacterial enzyme. This indicates that the protein-protein interactions necessary for proper proton coupling are essentially intact in the hybrid enzyme. The ATPase activities of the endogeneous and the hybrid enzymes were also very similar to each other, and Western blots indicated that the E. coli and spinach polypeptides were assembled into the FF complex to an approximately equal extent. The spinach subunit is thus a remarkably effective substitute for the native E. coli polypeptide. The failure of an earlier complementation attempt (Munn et al., 1991) can be attributed to use of an E. coli host in which residual assembly-defective F could block entry of chloroplast into the functional F complex. Gatenby and Rothstein (1986) found that a protein containing the first 365 amino acids of maize chloroplast CF fused to -galactosidase could be synthesized in E. coli and could associate with the bacterial membrane F complex. This fusion protein could not, however, assemble into an intact F complex (Gatenby and Rothstein, 1986).

Subunit Interactions and Cooperativity

Some recent studies have indicated that the amino terminus of the polypeptide is in contact with the subunit and that this contact is functionally important. First is the observation that Cys is buried within the CF complex and is accessible to modifying reagents only upon isolation of the subunit from the other CF subunits (Colvert et al., 1992). Second, the positions cognate to 63 and 64 in E. coli are sites of assembly mutants, some of which block assembly of both and subunits into the ATP synthase (Noumi et al., 1986; Miki et al., 1994a, 1994b). Third, these residues in E. coli are also proposed to lie at an interface, since they are recognized by a specific monoclonal antibody in the isolated but not the assembled (see Miki et al. (1994a, 1994b). Finally, this hypothesis is supported by the 2.8-Å crystal structure of beef heart mitochondria F (Abrahams et al., 1994). In the crystal structure, the amino-terminal region of the and subunits (mitochondrial amino acids 9-80, corresponding to 19-96 for CF ) is organized into a separate -barrel domain connected by a short hinge to the nucleotide-binding central domain. strand d from each subunit appears to contact strand a of the adjacent subunit, joining the hexameric ring of and subunits via their amino-terminal domains. Chloroplast residue 63 (47 in beef heart mitochondria) lies in strand d, and is thus implicated in subunit interaction by the structural evidence. These sequences, and the approximate location of the strands, are shown in Fig. 1B.

We examined several mutants at position 63 of the CF subunit. The conservative change (in terms of the size of the side chain) of Cys to Ala had little effect on the function of the subunit in the complementation assay. However, substitution of the bulkier side chains of Val or Trp reduced or blocked ATP synthesis in vivo. Indeed, ATP synthesis, as measured by succinate-dependent cell growth, was directly correlated with the volume of the residue at position 63, with the smallest residue being the most active, and the larger residues progressively less so. This result is consistent with Cys being located at a site of interaction between and subunits, and further demonstrates that this interaction is functionally important. Replacing Cys with the bulkier amino acids Val or Trp did not, however, prevent assembly of the subunit into the FF complex since washed membranes from both mutants contained approximately the same amount of subunit as did membranes from cells containing wild-type chloroplast or E. coli subunit. Moreover, ATPase activities of washed membranes from wild-type cells and from the C63A and C63W mutants were essentially identical. This result is further corroborated by the finding that these two mutant subunits, when over-expressed and refolded, bound nucleotides with close to the same affinity as native, wild-type spinach subunit.

The role of proton translocation through CF is to release bound product ATP (reviewed by Penefsky and Cross(1991)). On the other hand, ATP hydrolysis can drive reverse proton pumping, but may not be obligatorily coupled to it. The observation that the Cys Trp replacement appears specifically to block ATP synthesis but not ATP hydrolysis is particularly intriguing as it suggests that the bulky side chain interferes with cooperative subunit-subunit interactions required for coupling proton translocation to ATP synthesis but not with those involved in the catalytic cycle for ATP hydrolysis (reviewed by Penefsky and Cross(1991)). A complementary observation with E. coli F (Miki et al., 1994a) supports the independence of proton coupling and ATPase activity. A mutation from Glu to Asn at position -41 (cognate to CF -63) has 100% wild-type ATP synthase activity (judged by succinate-dependent growth), but only 4% wild-type ATPase activity.

The phenotype of the CF mutant -C63W indicates that specific subdomains of the interface may couple proton movement to ATP synthesis. Functional communication between Cys and the catalytic site is suggested by the following evidence. We determined (Colvert et al., 1992) that Cys is 42 Å away from the nucleotide binding site on the subunit. More recently, we have demonstrated that binding of ADP (but not ATP) to isolated CF- induces a shift in the conformation or position of the amino-terminal domain, forcing residue 63 into a more hydrophobic and less solvent-accessible environment (Mills et al., 1995). This change is detected as an increase in fluorescence quantum yield and a decrease in acrylamide-induced fluorescence quenching of pyrenyl maleimide attached to Cys. Taken together, these observations present functional evidence for a critical role of the amino-terminal domain of the subunit surrounding Cys in mediating cooperative interactions between the and subunits of CF, which are essential for ATP synthesis.

Functional Alterations in Hybrid ATP Synthase

We showed earlier (Richter et al., 1986) that reconstitution of the spinach chloroplast subunit into -less R. rubrum F resulted in a hybrid enzyme that was fully sensitive to the cyclic tetrapeptide tentoxin. Since tentoxin is a specific noncompetitive inhibitor of chloroplast F, this result meant that the polypeptide had conferred tentoxin sensitivity upon the hybrid enzyme. A recent report (Avni et al., 1992) suggested that sensitivity to tentoxin is determined by the nature of the amino acid residue at position 83 near the amino terminus of the subunit. Chloroplast residues 63-91 are identical (except for conservative changes at positions 77 and 83) in all sequenced tobacco species and in spinach. Tentoxin-resistant tobacco species contain Glu at residue 83, whereas tentoxin-sensitive tobacco and spinach possess Asp at this position (Fig. 1B) (Avni et al., 1992). Furthermore, conversion of Chlamydomonas reinhardtii residues 74-91 to the corresponding tobacco sequence produced algae that were tentoxin-resistant if residue 83 was Glu and tentoxin-sensitive if it was Asp (Avni et al., 1992). This result implied that Asp alone could direct binding of tentoxin. Wild-type E. coli F, on the other hand, is completely insensitive to tentoxin (Table 2), even though its subunit has an aspartate residue at the position (residue 60) corresponding to chloroplast position 83. Our results, furthermore, demonstrate that introduction of the tentoxin-sensitive spinach subunit into E. coli does not confer tentoxin-sensitivity to the transformed bacteria. Thus Asp alone is insufficient to determine tentoxin sensitivity. Instead, tentoxin binding probably requires a unique conformation of F resulting from an interaction between the amino terminus of the subunit and some other part or parts of the enzyme. We found recently (Hu et al., 1993) that tentoxin inhibits cooperative interactions between different nucleotide binding sites on CF and suggested that the toxin binds at an interface between the and subunits. However, residue 83, the putative tentoxin interaction site, is located at the top of strand e in the crystal structure of mitochondrial F (Abrahams et al., 1994) (see Fig. 1B), and may not be directly involved in subunit interactions. Binding of a molecule the size of tentoxin (packed volume = 426 Å; d 10 Å) at this site might well distort the interface, leading to loss of catalytic cooperativity. A more detailed hypothesis concerning the mechanism of tentoxin interaction can be formulated following publication of the atomic coordinates of the mitochondrial F crystal structure.

Conclusion

We have demonstrated that the chloroplast CF gene can be expressed in vivo in E. coli cells and assembled along with other E. coli FF subunits to form an active hybrid ATP synthase complex. In addition to allowing direct study of interactions between heterologous F subunits, the genetic complementation system will be of value in screening mutant spinach chloroplast subunits for their ability to fold correctly and to assemble into functional ATP synthase enzymes. Using this system, we have presented evidence consistent with the hypothesis that select amino-terminal subdomains of the subunit are engaged in important interactions that may play a critical role in the coupling of proton translocation to ATP synthesis but which are not essential for ATP hydrolysis.


FOOTNOTES

*
This work was supported by grants from the National Science Foundation (DMB-8805048 and OSR-9255223) and the United States Department of Agriculture (93-37306-9633) and from the University of Kansas (Biomedical Research Fund 4932, 4939). 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.

The nucleotide sequence(s) reported in this paper has been submitted to the GenBank®/EMBL Data Bank with accession number(s) U23082[GenBank® Link].

§
To whom correspondence should be addressed: University of Kansas, Dept. of Biochemistry, 2045 Haworth Hall, Lawrence, KS 66045-2106. Tel.: 913-864-3939; Fax: 913-864-5321; pgegen{at}kuhub.cc.ukans.edu

The abbreviations used are: CF, chloroplast coupling factor 0; CF, chloroplast coupling factor 1; LB medium, Luria-Bertani bacterial growth medium; TNP-ADP, 2`(3`)-O-(2,4,6-trinitrophenyl)adenosine 5`-diphosphate.


ACKNOWLEDGEMENTS

We thank Dr. Alan Senior for providing the E. coli deletion strain JP17 and plasmid pRPG31. The University of Kansas Biochemical Research Service Facility provided oligonucleotide synthesis and DNA sequencing services. We also thank Jing Ou for sequence confirmation of mutant plasmids and members of the Richter and Gegenheimer labs for helpful discussions.


REFERENCES
  1. Abrahams, J. P., Leslie, A. G. W., Lutter, R., and Walker, J. E.(1994)Nature 370, 621-628 [CrossRef][Medline] [Order article via Infotrieve]
  2. Avni, A., Anderson, J. D., Holland, N., Rochaix, J.-D., Gromet-Elhanan, Z., and Edelman, M. (1992)Science 257, 1245-1247 [Medline] [Order article via Infotrieve]
  3. Balbas, P., Soberon, X., Bolivar, F., and Rodriguez, R. L. (1988) in Vectors (Rodriguez, R. L., and Denhardt, D. T., eds) pp. 5-41, Butterworths, Boston, MA
  4. Boekema, E. J., and Bottcher, B.(1992)Biochim. Biophys. Acta 1098, 131-143 [Medline] [Order article via Infotrieve]
  5. Bordo, D., and Argos, P. (1991)J. Mol. Biol. 217, 721-729 [Medline] [Order article via Infotrieve]
  6. Boyer, P. D.(1993) Biochim. Biophys. Acta 1140, 215-250 [Medline] [Order article via Infotrieve]
  7. Burkovski, A., Lill, H., and Engelbrecht, S.(1994)Biochim. Biophys. Acta 1186, 243-246 [CrossRef][Medline] [Order article via Infotrieve]
  8. Chen, Z., Wu, I., Richter, M. L., and Gegenheimer, P. A.(1992)FEBS Lett. 298, 69-73 [CrossRef][Medline] [Order article via Infotrieve]
  9. Colvert, K. K., Mills, D. A., and Richter, M. L.(1992)Biochemistry 31, 3930-3935 [Medline] [Order article via Infotrieve]
  10. Engelbrecht, S., Lill, H., and Junge, W.(1986)Eur. J. Biochem. 160, 635-643 [Abstract]
  11. Gatenby, A. A., and Rothstein, S. J.(1986)Gene (Amst.)41,241-247 [Medline] [Order article via Infotrieve]
  12. Gibson, F., Cox, G. B., Downie, J. A., and Radik, J.(1977)Biochem. J. 164, 193-198 [Medline] [Order article via Infotrieve]
  13. Gunsalus, R. P., Brusilow, W. S. A., and Simoni, R. D.(1982)Proc. Natl. Acad. Sci. U. S. A. 79, 320-324 [Abstract]
  14. Hanahan, D. (1985) in Cloning: A Practical Approach (Glover, D. M., ed) Vol. I, pp. 109-135, IRL Press, Oxford, United Kingdom
  15. Hawthorne, C. A., and Brusilow, W. S. A.(1986)J. Biol. Chem. 261, 5245-5248 [Abstract/Free Full Text]
  16. Hu, N., Mills, D. A., Huchzermeyer, B., and Richter, M. L.(1993)J. Biol. Chem. 268, 8536-8540 [Abstract/Free Full Text]
  17. Lee, R. S-F., Pagan, J., Wilke-Mounts, S., and Senior, A. E.(1991) Biochemistry 30, 6842-6847 [Medline] [Order article via Infotrieve]
  18. Lill, H., Burkovski, A., Altendorf, K., Junge, W., and Engelbrecht, S.(1993) Biochim. Biophys. Acta 1144, 278-284 [Medline] [Order article via Infotrieve]
  19. Miki, J., Tsugumi, S., and Kanazawa, H.(1994a)Arch. Biochem. Biophys. 312, 317-325 [CrossRef][Medline] [Order article via Infotrieve]
  20. Miki, J., Kusuki, H., Tsugumi, S., and Kanazawa, H.(1994b)J. Biol. Chem. 269, 4227-4232 [Abstract/Free Full Text]
  21. Mills, D. A., and Richter, M. L.(1991)J. Biol. Chem. 266, 7440-7444 [Abstract/Free Full Text]
  22. Mills, D. A., Siebold, S. A., Squier, T. C., and Richter, M. L.(1995) Biochemistry 34, 6100-6108 [Medline] [Order article via Infotrieve]
  23. Munn, A. L., Whitfeld, P. R., Bottomley, W., Hudson, G. S., Jans, D. A., Gibson, F., and Cox, G. B.(1991)Biochim. Biophys. Acta 1060, 82-88 [Medline] [Order article via Infotrieve]
  24. Musier, K., and Hammes, G. G.(1987)Biochemistry 26, 5982-5988 [Medline] [Order article via Infotrieve]
  25. Nalin, C. M., and Nelson, N.(1987)Curr. Top. Bioenerg. 15, 273-294
  26. Noumi, T., Oka, N., Kanazawa, H., and Futai, M.(1986)J. Biol. Chem. 261, 7070-7075 [Abstract/Free Full Text]
  27. Olins, P. O., and Rangwala, S. H.(1990)Methods Enzymol. 185, 115-119 [Medline] [Order article via Infotrieve]
  28. Penefsky, H. S., and Cross, R. L.(1991)Adv. Enzymol. Relat. Areas Mol. Biol. 64, 173-214 [Medline] [Order article via Infotrieve]
  29. Reznikoff, W. S., and McClure, W. R. (1986) in Maximizing Gene Expression (Reznikoff, W. S., and Gold, L., eds) pp. 1-33, Butterworths, Boston, MA
  30. Richards, F. M. (1974)J. Mol. Biol. 82, 1-14 [Medline] [Order article via Infotrieve]
  31. Richter, M. L., Gromet-Elhanan, Z., and McCarty, R. E.(1986)J. Biol. Chem. 261, 12109-12113 [Abstract/Free Full Text]
  32. Rosenberg, A. H., Lade, B. N., Chui, D.-S., Dunn, J. J., and Studier, F. W.(1987) Gene(Amst.)56,125-135 [CrossRef][Medline] [Order article via Infotrieve]
  33. Rost, B., and Sander, C. (1993)Proc. Natl. Acad. Sci. U. S. A. 90, 7558-7562 [Abstract/Free Full Text]
  34. Schumann, J., Richter, M. L., and McCarty, R. E.(1985)J. Biol. Chem. 260, 11817-11823 [Abstract/Free Full Text]
  35. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk, D. C. (1985) Anal. Biochem.150, 76-85; Correction (1987) [Medline] [Order article via Infotrieve] Anal. Biochem.163,279
  36. Stormo, G. D., Schneider, T. D., Gold, L., and Ehrenfeucht, A.(1982) Nucleic Acids Res. 10, 2997-3011 [Abstract]
  37. Studier, F. W., Rosenberg, A. H., Dunn, J. J., and Dubendorff, J. W.(1990) Methods Enzymol. 185, 60-89 [Medline] [Order article via Infotrieve]
  38. Taussky, H. H., and Shorr, E.(1953)J. Biol. Chem. 202, 675-678 [Free Full Text]
  39. Walker, J. E., Fearnley, I. M., Gay, N. J., Gibson, B. W., Northrop, F. D., Powell, S. J., Runswick, M., Saraste, M., and Tybulewicz, V. L.(1985) J. Mol. Biol. 184, 677-701 [Medline] [Order article via Infotrieve]
  40. Zurawski, G., Bottomley, W., and Whitfeld, P. R.(1982)Proc. Natl. Acad. Sci. U. S. A. 79, 6260-6264 [Abstract]

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