MOLECULAR CLONING, INSERTIONAL INACTIVATION AND
CHARACTERIZATION OF THE CYANATE LYASE GENE FROM THE
CYANOBACTERIUM SYNECHOCOCCUS PCC 7942
Farid Jalali
A thesis submitted in conformity with the requirements for the degree of Master of Science Graduate Department of Botany University of Toronto
Q Copyright by Farid Jalali (1997) nbqummbtuvI.J ut IU -V~Y.YI.IV. IY Y. Bibliographic Services services bibliographiques 395 Wellington Street 395, rue Wellington Ottawa ON K1A ON4 Ottawa ON K1A ON4 Canada Cana& Your file Votre rtiference
Our fi& Notre réldrence
The author has granted a non- L'auteur a accordé une licence non exclusive licence allowing the exclusive permettant a la National Libraty of Canada to Bibliothèque nationale du Canada de reproduce, loan, distribute or sell reproduire, prêter, distribuer ou copies of this thesis in microform, vendre des copies de cette thèse sous paper or electronic formats. la fome de microfiche/nlm, de reproduction sur papier ou sur format électronique.
The author retains ownership of the L'auteur conserve la propriété du copyright in this thesis. Neither the droit d'auteur qui protège cette thèse. thesis nor substantial extracts fiom it Ni la thèse ni des extraits substantiels may be printed or othemise de celle-ci ne doivent être imprimés reproduced without the author's ou autrement reproduits sans son permission. autorisation. MOLECULAR CLONING, INSERTIONAL INACTIVATION AND CHARACTERIZATION OF THE CYANATE LYASE GENE FROM THE CYANOBACTERIUM SYNECHOCOCCUS PCC 7942
Farid Jalali, Master of Science 1997. Department of Botany, University of Toronto.
A 4.45 kb region of genomic DNA was isolated from the cyanobacterium Synechococcus PCC 7942 by means of screening subgenomic plasmid and phage libraries. Nucleotide sequence analysis of this region identified four open reading frames. ORF146 is 441 bp in length, coding for a protein of 146 amino acids that had 43% sequence identity to cynS from Escherichia coli The deduced amino acid sequence of the three open reading frames upstream of ORFI 46 showed significant sequence identity to proteins that are members of ABC transport systems. lnsertional inactivation of ORF146 using an antibiotic resistance gene cassette produced a strain (CS1) that was unable to metabotize cyanate. In particular, the CS1 strain was unable to support cyanate-dependent O2 evolution or chlorophyll g fluorescence quenching and did not produce CO2when presented with cyanate. lnsertional inactivation of the first gene of the putative ABC transport group (ORF440) yielded a strain (CAI) that exhibited a phenotype similar to that of CS1. Northern hybridization analysis identified two ORF146 transcript, one that was -0.5 kb and a larger transcript of -3.9 kb. These results provide molecular and physiological evidence for the identification of the cyanate lyase gene (cynS) from the cyanobacterium Synechococcus PCC 7942. Furthermore, these results indicate that the four genes identified form an operon. I have had the privilege of being Dr. George Espie's student for 3 years. I am truly thankful for his constant support and encouragement in every aspect of my work.
Over the years he has become more than my supervisor. I consider him a good friend and someone whom I regard with the greatest esteem. I would also like to thank my lab mates for their support and comedic interludes.
iii Page
ABSTRACT...... ii
..- ACKNOWLEDGEMENTS ...... ~ri
TABLE OF CONTENTS ...... iv
LIST OF FIGURES ...... viii
INTRODUCTION ...... 1 lnorganic Carbon Transport ...... 2
Use of Structural Analogs to Study CO, Transport ...... 5
Photosynthetic Metabolism of Cyanate in Synechococcus UTEX 625 ...... 7
Escherichia coli Cyanase ...... 10
Research Objectives ...... 15 Bacterial Strains and Growth Conditions ...... -17
E. coli Strains and Growth Conditions ...... 17
Growth Conditions for Synechococcus PCC 7942 ...... 17
Nucleic Acid Isolation ...... 18
Isolation of Genomic DNA from Escherichia coli KI2 ...... 18
Isolation of Genomic DNA from Synechococcus PCC 7942 ...... 19
Isolation of Plasmid DNA from Bacterial Cells ...... 20
Isolation of RNA from Synechococcus PCC 7942 ...... 26
DNA Manipulations ...... 22
PCR Amplification of Genomic DNA ...... 22
Restriction Digests and Agarose Gel Electrophoresis of DNA ...... 23
Gel Purification and Radiolabelling of DNA ...... 23
Southern Blotting and Hybridization ...... 24
Construction and Screening of Plasmid Libraries ...... 25
Construction and Screening of Lambda Phage Library ...... 27
Insertional Inactivations ...... 29
RNA Gels and Northern Analysis ...... 31
Measurement of Photosynthetic O, Evolution ...... 33
Fluorometry ...... 34
Mass Spectrometry...... 35 RESULTS ...... 38
Construction of Subgenomic Libraries and Isolation of a Genomic Region ...38
Sequence Analysis of pBH22. pE56 and pK50 ...... 44
ORF146 ...... 49
ORF289 ...... 51
ORF263 ...... 52
ORF440 ...... 56
Insertional Inactivation of ORF146 and ORF440 ...... 58
Cyanate Dependent O, Evolution and Fluorescence Quenching in WT and CS1 Cells of Synechococcus PCC 7942 ...... 67
Mass Spectrometric Analysis of Synechococcus PCC 7942 W and CS1 .....71
Cyanate-Dependent O, Evolution and Fluorescence Quenching in WT and CA1 Cells of Synechococcus PCC 7942 ...... 74
Cyanate Transport in WT, CSI, and CA1 Cells of
SynechococcusPCC7942 ...... 78
Northern Analysis of RNA Transcript Size From
WT, CS1 and CA1 Cells of Synechococcus PCC 7942 ...... -79 cynS is Responsible for Cyanate Metabolism in
SynechococcusPCC7942 ...... 82 cynS is Expressed as Part of an Operon ...... 85
Cyanate Uptake in Synechococcus PCC 7942 ...... -88
Physiological Significance of Cyanate Lyase in Synechococcus PCC 7942 ...93
Future Research Objectives ...... 96
Transcript Analysis ...... 96
Cyanate Transport ...... -97
Physiological Significance of Cyanate Lyase in
SynechococcusPCC7942 ...... 99
Concluding Remarks ...... 99
REFERENCES ...... 101 Page
Figure 1. Southern hybridization analysis of restriction enzyme digested genomic DNA from Synechococcus PCC 7942 ......
Figure 2. Organization of the genomic region isolated from Synechococcus
PCC7942 ......
Figure 3. Nucleotide and deduced amino acid sequence of the 4453 bp region identified from Synechococcus PCC 7942 ...... 45
Figure 4. Alignment of the deduced amino acid sequence of ORFI 46 with that of CynS from E. coli, and that of ORF289 with that of NrtD from
SynechococcusPCC7942 ......
Figure 5. Alignrnent of the deduced amino acid sequence of ORF263 with that of NrtB from Synechococcus PCC 7942 ......
Figure 6. Hydropathy profile of the deduced amino acid sequence of
ORF263 ......
viii that of NrtA f rom Synechococcus PCC 7942 ...... 57
Figure 8. Cloning strategy to generate plasmid pXH::ORF146 for
insertional inactivation of ORF146 from Synechococcus PCC 7942 ...... 59
Figure 9. Cloning strategy for construction of plasmid pP12::ORF440 to insertionally inactivate ORF440 from Synechococcus PCC 7942 ...... 61
Figure 10. Southern hybridization analysis of Synechococcus
PCC 7942 mutants, CS1 and CA1 ...... 64
Figure 11. Ci- and cyanate-dependent fluorescence quenching and 0, evolution in wild type and CS1 cells of Synechococcus PCC 7942 ...... 69
Figure 12. Mass spectrometric analysis of cyanate-dependent CO, eff lux from wild type and CS1 cells of Synechococcus PCC 7942 ...... 72
Figure 13. Ci and cyanate-dependent fluorescence quenching and 0, evolution from wild type and CA1 cells of Synechococcus PCC 7942 ...... 75 Y * * I wild type, CS1 and CA1 cells of Synechococcus PCC 7942 ...... -80 ABC, ATP-binding cassette; BME, bmercaptoethanol; BTP, 1.3- bis[tris(hydroxymethyl)-methylamino] propane; Ci, inorganic carbon; CA, carbonic anhydrase; CA1 , Synechococcus PCC 7942 cynA insertional inactivant; CP, carbamy l phosphate; CPase, carbarnyl phosphate synthetase; CS 1, Synechococcus PCC 7942 cynS insertional inactivant; CCM,carbon concentrating mechanism; COS, carbon oxysulf ide; CCCP, 3-chloro-carbonylcyanidephenylhydrazone; CTAB, hexadecyltrimethylammonium bromide; ; DCMU, 3-(,4-dichlorophenyl)-l , 1- dimethylurea; DDBJ, DNA Data Bank of Japan; DEPC. diethyl pyrocarbonate; DES, diethylstilbestrol; DNA, deoxyribonucleic acid; dNTP, deoxynucleotide triphosphate;
EDTA, ethylenediaminetetraacetic acid; EMBL, European Molecular Biology
Laboratories; EZ, ethoxyzolamide; F,, minimal fluorescence yield, Fmmaximum fluorescence yield; F,', variable fluorescence yield; FGRB, formaldehyde gel running buffer; LB, Lauria-Bertani; MOPS, 3-(N-morpho1ino)propane sulfonic acid; OCTase, ornithine carbamyl transferase; OD, optical density; ODU, optical density unit; PCC,
Pasteur Culture Collection; PCR, polymerase chain reaction; PDB, Protein Data Bank;
PPFD, photosynthetic photon flux density; RNA, ribonucleic acid; RNase, ribonuclease;
RuBP, ribulose 1,5-bisphosphate; Rubisco, ribulose 1,5-bisphosphate carboxylase- oxygenase; SA, specific activity; SD, standard deviation; SDS, sodium dodecyl sulfate;
Tris, Tris (hydroxymethyl) aminomethane; UTEX, University of Texas Culture
Collection; WT, wild type. INTRODUCTION
Cyanobacteria are prokaryotic organisms capable of oxygenic photosynthesis.
Photosynthetic carbon reduction proceeds via the reductive pentose phosphate pathway (Calvin cycle), which uses ribulose 1,5-bisphosphate carboxylase-oxygenase
(Rubisco) as the primary carbon fixing enzyme (Andrews and Lorimer, 1987). Rubisco uses CO, to catalyze the carboxylation of ribulose 1,5-bisphosphate (RuBP), resulting in the formation of two molecules of 3-phosphoglyceric acid. The carboxylation of RuBP by Rubisco is a remarkably inefficient reaction (Km (CO,). 300 PM. Kat 6.5 S-'), a problem that is compounded by Rubisco's ability to use 0, as an alternative and com petitive substrate. Rubisco's oxygenase activity is inherently wasteful since CO, is produced from glycolic acid metabolism and reductive energy is dissipated.
Cyanobacteria are able to circumvent these two problems, Rubisco's low affinity for
CO, and cornpetition by 4, by employing a mechanism that concentrates CO, at the active site of Rubisco. Two prominent components of the "CO, concentrating mechanism" (CCM) in cyanobacteria are inorganic carbon (Ci) transport systems and carboxysomes. Ci transport facilitates the uptake and concentration of Ci within cells, such that the intracellular concentration is greatly in excess of the extracellular concentration. Carboxysomes allow for the cornpartmentalkation of Rubisco and in turn may reduce its availability to 4, which would otherwise compete with CO, for substrate binding sites and lead to an overall decrease in photosynthetic output (Price and
Badger, 1991). Regardless of whether or not carboxysomes protect Rubisco from O,, the high level of CO, generated within the cells will compete much more effectively with
0, and thus reduce the level of the oxygenase reaction.
lnorganic Carbon Transport
The CCM of cyanobacteria serves to raise the intracellular concentration of CO, far in excess of that in extracellular environments. Convincing evidence exists for the
operation of distinct transport mechanisms that facilitate the uptake of both CO, and
HCO, (Badger 1987; Espie et al., 1991b; Miller et al., 1991 ). Ci transport is a light-
dependent process that is inhibited by inhibitors of energy metabolism such as DCMU,
CCCP, and DES (Espie et al., 1988; Miller et al., 1988; Miller et al., 1990; Shelp and
Canvin, 1984). Ci transport ability is significantly influenced by the Ci concentration
present during growth. Cells grown in the presence of high levels of Ci preferentially transport CO, (Miller and Canvin, 1987b), whereas growth at low concentrations of Ci
(in air) results in the induction of HCOj transport capability (Badger and Gallagher,
1987). Regardless of which Ci species is transported, the net result is the formation of
a large intracellular Ci pool. This pool is the immediate source of CO, for
photosynthesis and the rate of photosynthetic CO, fixation is in part determined by the
size of the pool (Kaplan et al., 1980; Espie and Kandasamy, 1992)
Initial evidence for active CO, transport was provided by Badger and Andrews
(1982) for a marine species of Synechococcus. Miller et al. (1988), using mass
spectrometry, demonstrated that the aquatic cyanobacterium Synechococcus UTEX QJ
625 was capable of depleting the extracellular medium of CO,, reducing it to near zero.
That the uptake of CO, was not dependent solely on CO, fixation was shown in
experiments in which fixation was inhibited by using iodoacetamide and glycoaldehyde
(Miller et al., 1988; Miller and Canvin, 1989). The use of these two inhibitors did not
effect the ability of Synechococcus UTEX 625 to deplete the extracellular medium of
CO, indicating that the observed rates of CO, transport were not solely a consequence
of CO, fixation.
Mass spectrometty is a convenient means of demonstrating the depletion of CO2 from the extracellular environment by cyanobacteria (Miller, 1990). Miller et al. (1988)
showed that the extracellular [COJdropped to near zero in spite of the fact that a large
amount of HCOj was still present in the medium. The depletion of CO2 to such an
extent thus caused the CO,-HCO; system to be in a state of chemical disequilibrium.
That this occurs, by its very nature, means that energy must be put into the system and
thus CO, uptake must be an active process (Miller et al., 1988). The fact that a
disequilibrium is established suggests an active process for the uptake of CO,. Further
evidence in support of an active transport mechanism comes from the fact that CO, is
transported against its concentration gradient which is estimated to be at least 5000 to 1 (Miller et al., 1991). That CO, transport requires energy input cornes from
observations which show a dependence on light and inhibition by darkness (Espie et
aL, 1988). Furtherrnore, operation of the CO2transport system is inhibited by inhibitors
of energy metabolism. Its energy requirement and saturation kinetics (Espie et al.,
1991) indicate that CO2transport may be a carrier mediated process in cyanobacteria. L+
However, positive identification of proteins or genes involved in the active transport of
CO, remain to be made.
Miller and Colman (1980a, b) provided direct evidence for HCO; transport in
Coccochloris peniocystis. Under alkaline conditions they observed rates of photosynthesis and HCO; transport that could not be supported by the spontaneous dehydration of HCO; to CO,. They also demonstrated that a majority of Ci uptake during steady state photosynthesis involved HCO; transport. HCO; transport in cyanobacteria is an active process, considering the fact that HCO,' is transported against a membrane potential of approximately -120 mV (Miller and Colman, 1980b;
Ritchie et al., 1991). The effects of inhibitors of energy metabolism on intracellular accumulation of HCO; provides further evidence for active HCO; transport (Miller and
Colman. 1980b). Whereas CO, transport appears to be constitutive, HCO; transport seems to be inducible. Depending on the concentration of Ci experienced during growth. Synechococcus UTEX 625 can transport HCOi via two modes (Espie and
Canvin, 1987; Espie and Kandasamy, 1992; McKay et al., 1993; Espie and
Kandasamy, 1994). These two modes differ with respect to their requirement for Na+.
Air grown cells of Synechococcus UTEX 625 transport HCO, via a Na+- dependent mechanisrn that requires between 25-40 mM extracellular Na' for maximum activity.
Standing culture cells (low Ci grown cells) do not have this requirement for Na+ and transport HCO, via a Na+-independent mechanism.
A second component of the CCM is carboxysomes. Carboxysomes are proteinaceous poiyhedral-shaped structures that are found in many autotrophic prokaryotes (Codd and Marsden, 1984). They are an important feature of the CCM since they have been found to house Rubisco (Coleman et al., 1982; Codd, 1988;
McKay et al., 1993). The protein coat of the carboxysome is thought to serve as a barrier to gases, preventing them frorn diffusing in or out of the carboxysome (Reinhold et al., 1989; Reinhold et al., 1991). This presumed permeability barrier in turn reduces diffusion of O, into the carboxysome, which subsequently decreases corn petition for binding sites at Rubisco between 0,and CO,. Although the diffusion barrier prevents passage of O, into the carboxysome, its apparent that HCOi, the predominant intracellular Ci species, can pass into the carboxysome (Reinhold et aL, 1989). lnside the carboxysome, HCO, must be converted to CO, for it to be used by Rubisco for the carboxylation of RuBP. Since the spontaneous conversion of HCO,' to CO, under physiological conditions is slow, enzyme-mediated conversion is necessary (Reinhold et al., 1989). The presence of carboxysomal carbonic anhydrase (CA) facilitates the rapid conversion of HC03-to CO,, which in turn can be fixed by Rubisco (Badger and
Price, 1989; Yu etaL, 1992). Since the protein coat is impermeable to gases, the CO, produced by CA accumulates within the carboxysome, and in turn around the active site of Rubisco. In this way, cyanobacteria are able to support rapid rates of photosynthetic carbon fixation.
Use of Structural Analogs to Study CO, Transport
Considerable debate has occurred in the literature as to whether or not CO, is an actual substrate of a protein carrier, since it easily permeates lipid membranes. The use of isotopic and structural analogs has gone a long way in resolving this question.
Espie et al. (1991a) showed that 13C02uptake was inhibited by 12C02under conditions that precluded HCO,' transport (due to an absence of Na'). When H13CO; transport was perrnitted, 12C02did not reduce HI3CO; transport, but continued to effectively inhibit %O, uptake. The inhibition of I3CO, uptake by I2CO2is competitive since increasing concentrations of 12C0, result in increased inhibition.
Carbon oxysulf ide (O=C=S, COS), an isoelectronic structural analog of CO,, has also been shown to effectively inhibit CO, transport without significantly impairing Na+- dependent HCO; transport (Ogawa and Togasaki, 1988; Miller et al., 1989; Badger and Price, 1990). Kinetic analysis indicated that COS was a competitive inhibitor of
CO, transport and thus interacted at a common active site. Similarly, cyanate (O=C=N',
OCN] another structural analog has also been shown to inhibit CO2 transport but not
HCO,' transport (Espie and Tong, unpublished). Surprisingly, both COS and OCN' are metabolized by Synechococcus UTEX 625 and PCC 7942 (Miller et al., 1989; Miller and Espie, 1994). COS is converted to CO2 and H2S, possibly by CA. Of note is the fact that H2S is also a selective inhibitor of CO, transport (Espie et aL, 1989). Interestingly,
COS, H,S and OCN' are al1 known inhibitors of CA, while CO, is a known substrate.
Thus, it seems likely that the CO2 transport system recognizes CO, through a membrane bound CA-like activity . This is further supported by the fact that the CA inhibitor ethoxyzolamide (EZ), which binds to the catalytically active Zn+ of CA, also inhibits CO, transport (Price and Badger, 1989; Tyrrell et al., 1996). Of direct concern f to my work is that OCN' is decomposed to CO, and NH, by cyanase in the
cyanobacteriurn Synechococcus UTEX 625 (Miller and Espie, 1994).
Photosynthetic Metabolism of Cyanate in Synechococcus UTEX 625
Cyanate is an anion that has been tested as an inhibitor of HCO; and CO, transport in Synechococcus UTEX 625 (Miller et al., 1991; Espie and Tong,
unpublished). Recently, Miller and Espie (1994) determined that rather than inhibit
photosynthesis, cyanate supported a rapid rate of photosynthetic O, evolution in the
cyanobacterium Synechococcus UTEX 625. Addition of 1 rnM cyanate to cells at the
CO, compensation point resulted in light-dependent O2 evolution at a rate of up to 188
prno102mg-' Ch1 h-'. This rate was equal to the maximum rate of photosynthesis at CO,
saturation. To support this rate of photosynthesis, cyanate decomposition to CO, would
have to occur at a rate equivalent to 94 nmol min'' per 1Og cells. In comparison, fully
induced E. coli cells decomposed cyanate at a maximum rate of 4.4 nmol min" per 1 0'
cells (Sung and Fuchs, 1989). Thus, Synechococcus UTEX 625 has a tremendous
capacity for cyanate metabolism.
Convincing evidence for cyanate decomposition was obtained using mass
spectrometry and crude cell extracts (Miller and Espie, 1994). When K013C~was
provided the extract evolved 13C labelled Ci. The immediate product of K013CN
decomposition was found to be 13C02,not H13C0,', since the addition of CA to the
lysate caused a drop in the 13C02signal. If H13C0,' was the immediate product of K013CN decomposition, an increase in the 13C02would be ewected, resulting from CA- mediated conversion of H13CO; to 13C02.However, 13C02production from K013CNwas stimulated by and dependent upon the concentration of KHCO, present in the lysate suggesting a role for HCO; in the decomposition of cyanate.
Further analysis indicated that Synechococcus UTEX 625 also produced NH, when provided with KOCN. These observation lead the authors to propose that
Synechococcus UTEX 625 possessed the enzyme cyanase (or cyanate lyase) which catalyzed the following reaction:
OCN' + 2H+ + HCO, + ZCO, + NH, where HCO; served as a recycling substrate, as suggested for E. coli cyanase by
Johnson and Anderson (1987). This proposal was supported by the observation that oxalate, a known inhibitor of E. colicyanase, blocked 13C02production from K013CN in Synechococcus UTEX 625 lysates. Fractionation studies showed that cyanase activity was absent in the periplasm, the carboxysomes and the thylakoid membranes, but that it was present in the soluble fraction of the cytosol (Miller and Espie, 1994).
That cyanase activity was observed from cells that were not pre-exposed to cyanate, indicates that it is expression is constitutive under the growth conditions they used
(Miller and Espie, 1994).
The addition of KO',CN to intact, illuminated Synechococcus UTEX 625 cells did not result in the appearance of CO, in the reaction medium (Miller and Espie,
1994). However, when the light was turned off a large efflux of 13C0, was observed.
The addition of COS,a CO,transport inhibitor, prior to KO'~CNalso resulted in a large light-dependent efflux of 13C02.Based on these observations Miller and Espie (1994) proposed the following scheme to account for cyanate-dependent 0, evolution.
Cyanate enters cells by a mechanism that remains to be determined. Once inside cyanate is decomposed to CO, and NH, by cyanase. HCO, for the reaction is provided by the CCM which builds up a large intracellular Ci pool. The CO, that is produced f rom cyanate has several fates. CO2 rnay be fixed by the Calvin cycle andor it rnay also leak out of the cell, in which case it rnay be recycled by the CO, transport system.
Ultimately, CO, that is generated from cyanate will be fixed leading to photosynthetic
4 evolution. NH, that is generated rnay be fixed via the nitrogen assimilatory pathway into metabolic compounds or it rnay leak out of the cell. It seems reasonable to assume and preliminary evidence suggests (Miller and Espie, unpublished) that cyanate rnay serve as the sole source of C and N for growth. However, these experiments are confounded by the spontaneous decomposition of cyanate to CO, and NH, in aqueous solutions (Allen and Jones, 1964). Only illuminated cells can metabolize cyanate.
Although the reason for this remains unknown, energy input at the level of cyanate uptake remains a possibility. The light requirement rnay also reflect energy input at the level of Ci transport, which provides cyanase with it's other essential substrate, HCO,.
The precise nature of the light requirement for cyanate metabolism remains to be determined. Escherichia coli Cyanase
Cyanase (EC 4.3.99.1), or cyanate lyase, is responsible for the bicarbonate- dependent decomposition of cyanate to carbon dioxide and ammonia in E. coli
(Anderson et al., 1990). It's activity was f irst described in Escherichia coli BI1 that was grown in the presence of potassium cyanate (Taussig, -1 960). Cyanase activity has since been identified in several strains of E. coli, Pseudomonas fluorescences NClB
11764 (Kunz and Nagappan, 1989), a species of Flavobacterium sp. (Guilloton and
Hargreaves, 1972) and most recently in the cyanobacterium Synechococcus UTEX 625
(Miller and Espie, 1994). With the exception of Synechococcus, cyanase activity is only detected when the cells are pre-exposed to cyanate in the growth medium, suggesting that cyanase activity was inducible. A physiological role for cyanase in E. coli has yet to be determined conclusively, but it seems likely that cyanase eliminates cyanate, which can have inhibitory effects on growth (Guilloton and Karst, 1987b; Kozliak et al.,
1995).
In E. coliK12, cyanase is coded for by cynS, a member of the cyn operon. The cyn operon also codes for carbonic anhydrase (cyn7) and a hydrophobic protein of unknown function (cynX) (Sung and Fuchs, 1988; Sung and Fuchs, 1989; Guilloton et al. 1992). The cynS gene was initially identified because of its linkage with the lac operon. Mutants with defects in the lac operon were found to have an increased sensitivity to cyanate and an inability to utilize cyanate as a sole source of nitrogen for growth (Guilloton and Karst, l987a; Sung et al., l987a). Sung et al. (1Q87b), using 11 these lac operon mutants, successfully isolated and overexpressed the cynS gene.
Although the deduced amino acid sequence of cynSfrom E. coli KI2 differed slightly from the actual amino acid sequence determined for E. co6 BI1 (Chin et al, 1983). its
physical and kinetic properties did not differ from those described by Anderson (1 980) and Anderson and Little (1986).
Transcription of the cyn operon yields two transcripts that share common 5' ends, but differ at the 3' end by approximately 1000 bp (Sung and Fuchs, 1988). The smaller codes for both cynS and cynT, whereas the longer includes cynX. ln vivo termination studies determined that approxirnately 75% of the transcripts were of the shorter variety, resulting in decreased expression of cynX (Sung et al., 1987b). The cyn operon has two transcription terminators. Use of T7 RNA polymerase (which does not recognize E. coli transcription terminators) results in increased expression of cynX, suggesting that the first terminator functions in downshifting expression of cynX by terminating its transcription prematurely. A function for this mechanism has yet to be determined.
Transcription of the cyn operon is regulated by CynR, which is a member of the
LysR family of regulatory proteins (Sung and Fuchs, 1992). The cynR gene is located upstrearn of the cyn operon, however, its transcription occurs in the opposite direction.
CynR is responsible for regulating transcription of the cyn operon, as well as itself. In the presence of cyanate it activates transcription of the cyn operon. lt has been suggested that CynR alternates between two functionally different conformations depending on the absence or presence of cyanate. In the absence of cyanate, CynR IL
binds at a regulatory site, preventing transcription of the cyn operon as well as itself.
Positive regulation of the cyn operon ensures that it is not expressed in the absence of cyanate, and negative autoregulation of cynR by CynR maintains its level at an approximately constant level.
Cyanase catalyzes the bicarbonate-dependent decomposition of cyanate to form two molecules of CO, according to the reaction (Johnson and Anderson, 1987):
OCN' + 2H' + HCO", + 2C0, + NH,
Considering the reaction catalyzed, cyanase has a rather complex quaternary structure. The native enzyme has a molecular weight of approximately 150 KDa, with each identical subunit having a molecular weight of approximately 17 KDa (Chin et al.,
1983). The native enzyme has been crystallized (Kim et a/. 1987), and preliminary X-
ray crystallographic data suggests a 5/2 symmetry, that is, a pentamer of dimers
(Otwinowsky et al, 1991). Inactive dimers, which are thought to be stabilized by a disulfide bond between the single cysteine residues of a subunit pair, associate to form an active decamer (Little and Anderson, 1987; Anderson et al., 1994). A search for active site residues resulted in the finding that derivitization or site directed mutagenesis of the single cysteine residue does not impair catalytic activity since decameric structure can be obtained (Little and Anderson, 1987; Anderson et a/.,1988;
Anderson et al., 1994). The decamers that are obtained by derivitization or site directed mutagenesis are catalytically active, however, their stability is effected dramat ically.
They undergo reversible dissociation to inactive dimers under conditions that weaken hydrophobie interactions, or when substrate or substrate analogs are absent (Anderson et al., 1994). Similar results are obtained when the single histidine residue of a subunit is replaced by an asparagine or tyrosine residue. Therefore, the single cysteine and histidine residues of a subunit are not required for catalytic activity, but are necessary for stabilizing decameric structure, which is a prerequisite for catalytic activity. They are also required for facilitating association of inactive dimers to give active decamers. The requirement of bicarbonate and/or azide (a structural analog of cyanate) for decamer formation from mutant dimers suggests that they, in some way, participate in stabilizing the decamer, perhaps by shifting an equilibrium in the favour of decamer, since the free dimer does not bind substrate. Inactive dirners associate to give active decamers, however, in this conformation they are not linked via disulfide bonds. Anderson et al.
(1 994) suggest that the sulfhydryl groups of a dimer are spatially close to each other and that formation of a decamer involves functionally significant conformational changes within the dimer. These changes are thought to prevent disulfide bond formation and are either required or affect catalytic activity.
The bicarbonate-dependent breakdown of cyanate yietds CO, and NH,. Initial kinetic analyses of the cyanase reaction was perf ormed by Anderson (1980) and more extensively by Anderson and Little (1986).The cyanase reaction proceeds via a rapid equilibrium random mechanism, where both cyanate and bicarbonate, exhibit competitive su bstrate inhibition (Anderson and Little, 1986). Su bstrate inhibition occurs when either substrate binds at the others binding site, resulting in the formation of a dead-end cornplex. The reaction of cyanate with bicarbonate does not immediately result in the formation of carbon dioxide and ammonia. The initial product of the reaction is carbamate which then spontaneously breaks down to carbon dioxide and ammonia. The reaction to form carbamate proceeds via the transient formation of a dianion intermediate. Support for the existence of a dianion intermediate comes from the fact that oxalate (a dianion) acts as a potent inhibitor of cyanase. Binding studies indicate that oxalate exhibits a stoichiometry of one binding site per two subunits, suggesting that it may occupy both sites at once Le. a dianion intermediate (Anderson et al., 1987).
The use of bicarbonate as a substrate for the cyanase-mediated decomposition of cyanate is interesting since it seems possible that cyanate could react directly with water, through carbamate, resulting in the production of one carbon dioxide and ammonia (Johnson and Anderson, 1987). Alternatively, bicarbonate may serve as an activator of cyanase. Anderson (1980) first suggested that bicarbonate is an active participant in the cyanase reaction, and that it may function as a recycling substrate.
Johnson and Anderson (1987) demonstrated that bicarbonate does serve as a recycling substrate, since half of the carbon dioxide that is produced is derived from bicarbonate.
A physiological role for cyanase in E. coli has as yet to determined. It has been suggested that cyanase is induced to remove cyanate before it inhibits growth
(Anderson et al., 1990). Although cyanate is not commonly found in nature it can arise from the spontaneous breakdown of urea in aqueous solutions (Hagel et al., 1971 ). It can also arise from the spontaneous breakdown of carbamoyl phosphate (Allen and
Jones, 1964). Cyanate is toxic since it can react with the functional groups of proteins. Cyanate can carbamylate protein amino groups in a reaction that is irreversible.
However its reaction with thiols, imidazoles, phenolic groups, and carboxylates is reversible (Stark, 1972). As a result of its reactive nature, cyanate is a known inhibitor of several enzymes. Cyanate reacts with enzymes, causing structural and/or functional alterations as a result of carbamylation. Notable examples of enzymes that are inhibited by the reaction of cyanate with a functional group involved in catalysis are carbamoyl phosphate synthase from E. coli (Anderson and Carlson, 1975), and ribulose 1,5-bisphosphate carboxylase/oxygenase from tobacco (Chollet and
Anderson, 1978). Cyanate is also regarded as a pseudohalide that can form corn plexes with transition metal ions (Forster and Goodgame, 1965). As a pseudohalide, cyanate mediates inhibition of enzyme activity after chelation of metal ions at an enzymes active site. Notable examples are carbonic anhydrase III from bovine skeletal muscle
(Engberg and Lindskog, 1984) and superoxide dism utase (Bertini et al., 198 1).
Research Objectives
Although cyanase (or cyanate lyase) is well characterized in E. coli, little is known about it and the rote it plays in other organisms. The novel interaction between cyanase and the photosynthetic apparatus of Synechococcus is almost completely unknown and suggests that cyanate metabolism may have a distinct, and different function and rnechanism in photosynthetic autotrophs. The physiological and molecular characteristics of the photosynthetic apparatus of Synechcococcus PCC 7942 are well characterized. Furthermore, Synechococcus PCC 7942 is easily manipulated using
basic recombinant DNA techniques. Given the fact that it is naturally transformable,
Synechococcus PCC 7942 provides an ideal system for investigating the role of
cyanase in the context of photosynthetic metabolism. The intention of this study,
therefore, was to identify gene(s) involved in cyanate metabolism and in particular the
gene for cyanate lyase in the cyanobacterium Synechococcus PCC 7942. To identify this gene, the cynS gene from E. coli will be used to probe the genome of
SynechococcusPCC 7942 for a cynS- homolog. Once identified and cloned, the
Synechococcus cynS gene will be insertionally inactivated to provide physiological
evidence for its involvement in cyanate metabolism. Cyanate has been used as an
inhibitor of CO, transport, however its effectiveness as an inhibitor is confounded by the fact that it is metabolized. The strain generated in this study will serve as a platform for the analysis of CO, transport using cyanate as an inhibitor. MATERIALS AND METHODS
Bacterial Strains and Growth Conditions
E. coli Strains and Growth Conditions. E. coli KI2 (kindly provided by Dr. Barbara
Funnel, University of Toronto) was maintained on Lauria-Bertani (LB) (Sambrook et a/., 1989) media with 1.4% [wlv] agar (plate cultures). For isolation of genomic DNA,
E. coliK12 was grown in liquid LB media at 37°C with shaking. E. coliXL1 Blue MRF'
(Stratagene) was maintained on LB plates supplemented with 12.5 pg mLml tetracycline.
E. coli SOLR cells were maintained on LB plates supplemented with 50 pg ml-' kanamycin. E. coli DH5a (Gibco-BRL) cells harbouring plasmids were maintained on
LB plates supplemented with 100 pg ml" ampicillin. For large scale isolation of plasmid DNA, E. coli DH5a cells were grown in LB liquid media with 100 pg ml'' ampicillin.
Growth Conditions for Synechococcus PCC 7942. Cells of the cyanobacterium
Synechococcus PCC 7942 were maintained photoautotrophically on 1.4% [w/v] agar plates containing BG-1 1 media at 30°C and light was provided at 20 pmol (photons) m'2s'' (PPFD). Standing culture cells (SCC) of Synechococcus PCC 7942 were grown in 50 mL of liquid BG-11 media at 30°C and the same illumination. Air-grown cells
(AGC) of Synechococcus PCC 7942 used for the isolation of genomic DNA, were grown in 1 L flasks containing 800 mL BG-1 1 medium and were supplied with air at a constant flow rate (540 mL min-'). The chlorophyll a concentration of cyanobacterial cultures used for various experiments was deterrnined using the method of MacKinney
(1941). A given volume of cells was centrifuged and the pellet extracted with methanol at -20°C. The concentration of chlorophyll a was determined spectrophotometrically at OD,,,.
Nucleic Acid Isolation
Isolation of Genomic DNA from E. coli K12. Genomic DNA from E. coli KI2 was isolated using a minipreparation technique outlined by Ausubel et al. (1 994). A 1.5 mL volume of overnight bacterial culture was centrifuged and the resulting bacterial pellet resuspended in 567 pL of TE buffer (10 mM Tris-HCI pH 7.5, 1 mM EDTA pH 8.0). To this mixture, 30 pL of 10% SDS and 3 pL of 20 mg ml" Proteinase K were added and the mixture incubated at 37°C. After 1 hour, 100 PL of 5 M NaCl was added. Following thorough mixing of the suspension, 80 pL of a CTABINaCI solution (10% CTAB in 0.7
M NaCI) was added, and the suspension incubated at 65OC for 10 minutes. An equal volume of chloroform/isoamyl alcohol (24:l) was added, the solution mixed, and centrifuged for 5 minutes at 12 000 x g. The aqueous phase was extracted again with an equal volume of phenol/chloroformlisoamyl alcohol (24:24:1), and centrifuged for another 5 minutes. The aqueous phase was recovered and nucleic acids were precipitated by adding 0.6 volumes of isopropanol. The DNA was pelleted by centrifugation, and washed with 70% ethanol. After centrifugation, the pellet was air dried and resuspended in an appropriate volume of sterile water. The quality of the
DNA obtained was determined by comparing absorbance at OD,, and OD,,, and by
restriction enzyme digestion and agarose gel electrophoresis .
Isolation of Genomic DNA from Synechococcus PCC 7942. Genomic DNA from
Synechococcus PCC 7942 was isolated essentially using the technique outlined by
Kuhlemeier and Van Arkel (1987) with some modifications. Cells that were air-grown
in 800 mL batch cultures were centrifuged at 4000 x gfor 10 minutes at 4°C. The pellet was washed once with SE (120 mM NaCI, 50 mM EDTA pH 8.0), centrifuged and
resuspended in lysis buffer (25% sucrose, 50 mM Tris-HCI, 100 mM EDTA pH 8.0) to
approximately 40 mL g-' wet weight. The resulting cell suspensions were transferred to 250 mL Erlenmeyer flasks (40 mL per flask) and incubated at 37°C for 1 hour with
shaking in the presence of 3 mg ml" lysozyme. After 1 hour, 14 mL (per 40 mL
suspension) of 10% SDS was added and the suspension incubated at 37°C for an additional 1 hour with shaking. To the shaking suspension, 6 mL of 5 M sodium
perchlorate was added. The suspensions were then transferred to Sorvall tubes and extracted with an equal volume of phenol/chloroform (1: 1) at 4°C with gentle shaking for 20 minutes. This mixture was centrifuged at 8000 x g for 10 minutes, and the aqueous phase extracted with chloroform at 4°C until the interphase was clear.
Subsequent to centrifugation, nucleic acids were precipitated at -20°C for 2 hours by the addition of 0.1 volumes of 3 M sodium acetate (pH 6.5) and 2.5 volumes of ice cold absolute ethanol. Nucleic acids were centrifuged at 12 000 x g for 10 minutes at 4°C and resuspended in 1.O mL of sterile water. The resulting nucleic acid solution was treated with RNase (75 pg mL1)for 1 hour at 37°C. RNase and residual proteins were removed by treatment with Proteinase K (50 pg L*') for 1 hour at 37OC. The DNA solution was phenol/chloroform extracted as described above and DNA was precipitated as described previously. The final DNA pellet was resuspended in sterile water. The concentration of DNA obtained was detemined using absorbance at 04,
(1 ODU= 50 pg ml-').
Isolation of Plasmid DNA from Bacterial Cells. To quickly determine the size of recombinant plasmids in E. coli DH5a or SOLR, a rapid cell disruption protocol
(Sambrook et al., 1989) was used with some modifications. A small quantity of bacterial cells was transferred from a plate to a microcentrifuge tube with a sterile toothpick. The bacterial cells were resuspended in 100 pL of a lysis solution containing 10 mM EDTA
(pH 8.0),0.2 N NaOH, 0.5% SDS and 20% sucrose. The resulting cell suspension was vortexed vigorously for 20 seconds, and then transferred to a 70°C water bath for 5 minutes. After the mixture was allowed to cool to room temperature, 1.5 PLof 4 M KCI and 0.5 PL of 0.4% bromophenol blue were added. The mixture was vortexed vigorously for 20 seconds and placed on ice for 5 minutes. The mixture was centrifuged at 12000 x g at 4°C for 3 minutes, and 50 pL of the supernatant was examined for plasrnid DNA using agarose gel electrophoresis.
To isolate large quantities of high purity plasmid DNA for nucleotide sequencing, cloning and other DNA manipulations, a plasmid preparation kit was used. The RPM AFS (BI0 101, Inc., Vista, Ca.) midiprep plasmid preparation kit was used in accordance with the manufacturer's instructions without modifications. The methodology is essentially that of alkaline lysis, however, pure plasmid DNA is isolated by the use of a glass DNA-binding matrix that gives very a high yield of supercoiled plasmid DNA.
Isolation of RNA from Synechococcus PCC 7942. Isolation of RNA from
Synechococcus PCC 7942 was carried using the methods outlined by Luque et al.
(1994) and Strommer et al. (1993) with modifications. Cyanobacterial cells (up to 1 gram, wet weight) were hawested at 8000 x gfor 5 minutes at 4°C. The cell pellet was washed once in 2.5 mlof 0.3 M sucrose and 10 mM sodium acetate (pH 4.7) and centrifuged at 8000 x g for 5 minutes at 4°C. The pellet was resuspended in 1 mL of
0.3 M sucrose, 10 mM sodium acetate, and 60 mM EDTA (pH 8.0). Liquid nitrogen was added to the cell suspension in a mortar and a porcelain pestle was used to grind the cells. Once the cells had been ground to a fine powder, 10 rnL of extraction buffer (4
M guanidinium isothiocyanate, 25 mM sodium citrate, 0.5% sarkosyl, and 0.1 M BME) was added directly to the ground cells in the mortar. The lysates were then transferred to 50 mL Sorvall centrifuge tubes. After vortexing for 30 seconds, 1 mL of 2 M sodium acetate (pH 4.7) was added. After vortexing the mixture for 30 seconds, 10 mL of buffer-saturated phenol (pH 4.5) was added, and the mixture was again vortexed for
30 seconds. Finally, 2 mL of chloroforrn was added and the mixture vortexed for 30 seconds. The resulting cell lysate was centrifuged at 5000 x g for 10 minutes at roorn temperature. The aqueous phase was removed and to it an equal volume of isopropanol was added. This mixture was incubated at -20°C for 1 hour, and then centrifuged at 10 000 x gfor 10 minutes at 4°C.The resulting pellet was resuspended in 500 pL of DEPC-treated water. Subsequently, an equal volume of 4 M LiCl was added and the mixture incubated on ice for 2.5 hours. The mixture was then centrifuged at 12 000 x gfor 5 minutes, and the resulting pellet washed two times with 70% ice cold ethanol. After the final centrifugation, the RNA pellet was resuspended in an appropriate volume of DEPC-treated water and the RNA quantified using spectrophotometry, where 1 ODU= 40 pg RNA ml".
DNA Manipulations
PCR Amplification of Genomic DNA. Two oligonucleotide primers, CYNSI and
CYNS2, were synthesized using the known sequence of cynS from E. coli K12.
Polymerase chain reaction (PCR) was performed essentially using the method outlined by Saiki et a/. (1990). Approximately 1 ng of E. coli KI 2 genomic DNA in 10 PL of sterile
H,O was mixed thoroughly with 10 pL of a 2 X PCR reaction mix containing 0.01% (wlv) gelatin, 10 mM Tris-HCL pH 8.3, 50 mM KCI, 1.5 rnM MgCI,, 200 uM of each dNTP, 1 pM of each primer and 1 unit of Thermus aquaticus (Taq) DNA polymerase (Perkin-
Elmer Cetus). Each PCR reaction was overlaid with 25 pL of sterile light mineral oil prior to placement in the thermal cycler. PCR amplification was perforrned using a DNA
Thermal Cycler 480 (Perkin-Elrner Cetus ). Standard PCR reactions were carried out 23 for 30 cycles, with each cycle made up of 3 steps: denaturation at 94°C for 1 minute, primer annealing at 50-57°C for 1 minute, and primer extension at 72°C for 1 minute.
After the completion of 30 cycles, there was an additional primer extension step for 10 minutes, followed by stoppage of reactions by incubation at 4°C.
Restriction Digests and Agarose Gel Electrophoresis of DNA. Restriction endonucleases purchased from New England Biolabs and Gibco-BRL were used to digest genomic and plasmid DNA, in accordance with the manufacturer's instructions.
Prior to electrophoresis, loading dye (0.25% [wlv] bromophenol blue, 0.25% [wlv] xylene cyanol FF, 15% [wlv] Ficoll Type 400 (Pharmacia)) was added to the restricted
DNA samples, that had been incubated at 65OC for 5 minutes before being placed on ice. Restricted and PCR amplified DNA samples were separated using 0.8% and 1.2%
[w/v] agarose gels, respectively, (unless otherwise specified) consisting of 1 X TAE
(0.04 M Tris-acetate, 1mM EDTA) running buffer. Electrophoresis was carried out for
2 hours at 60 V (unless otherwise specified), and gels were then stained with 0.5 JJ~ mL1 ethidium bromide and photographed using the Gel Doc 1000 lmaging System
(Bio-Rad).
Gel Purification and Radiolabelling of DNA. DNA fragments of interest were gel purified using the QlAEX II Gel Extraction Kit (QIAGEN) in accordance with the manufacturers instructions. Gel slices were cut out of ethidium bromide-stained gels under UV light. Gel-purified DNA fragments used for Southern, Northern, colony or L;L) plaque hybridizations were radiolabelled with [a-32P]dCTPusing an Oligolabelling Kit
(Pharmacia). Radiolabelled DNA was further purified using Bio-Gel P-10 (Bio-Rad) size exclusion chrornatography (Sambrook et a/., 1989).
Southern Blotting and Hybridization. Southern transfer of DNA from agarose gels ont0 Zeta-Probe (Bio-Rad) nylon blotting membrane was performed according to the instructions provided by the manufacturer. Gels were treated with a depurinating acid solution (0.25 M HCI) for 10 minutes with gentle shaking. The gels were then transferred to a denaturing solution (0.5 M NaOH, 1 M NaCI) for 30 minutes with gentle shaking, and subsequently treated with a neutralizing solution (0.5 M Tris-HCI pH 7.4,
3 M NaCI) for 30 minutes with gentle shaking. A sponge was placed in a flat bottom container, soaked with 10 X SSC (150 mM sodium citrate, 1.5 M NaCI), with additional
10 X SSC added to reach half way up the height of the sponge. Three pieces of
Whatman 3MM paper were placed in the middle of the sponge and soaked with 10 X
SSC. The gels were then placed on top of the paper, and the entire surface of the container was covered with plastic wrap. A window corresponding to the size of the gel was cut out of the plastic wrap. A piece of nylon membrane, pre-wetted in distilled water, followed by two additional pieces of pre-wetted 3MM paper were placed on the gel. Blotting was initiated by the addition of a 15 cm stack of paper towels and 1 plexiglass plate. Blotting was generally carried out for 12-18 hours.
After blotting, nylon membranes were gently rinsed with 2 X SSC and vacuum dried at 80°C for 30 minutes. Dried membranes were pre-hybridized in glass bottles with 6 X SSC, 0.5% SDS, and 5 X Denhardt's solution (O. 1% [wlv] BSA Fraction V, 1%
Ficoll Type 400, 0.1 % [wh] polyvinylpyrollidone) at 55-65°Cfor 2-4 hours in a Tek Star
Jr. hybridization oven (Bio/Can). When prehybridization was cornplete, the liquid content of the bottle was removed and replaced with hybridization solution (same as pre-hybridization with the addition of 10% [w/v] dextran sulfate and 100 ~g ml-' heat- denatured salmon sperm DNA). Heat-denatured radiolabelled probe DNA was added to the hybridization solution, and hybridization was allowed to proceed for 12-18 hours.
Membranes were washed twice with 2 X SSC at room temperature for 5 minutes, twice in 2 X SSC-1% SDS at the hybridization temperature for 30 minutes, and twice in 0.1
X SSC at room temperature for 10 minutes. Blots were placed on 3MM paper pre- wetted with 0.1% SSC, covered in plastic wrap and examined using autoradiography.
Blots were exposed to Kodak X-Omat AR imaging film in the presence of Cronex
(Dupont) intensifying screens at -70°C.
Construction and Screening of Plasmid Libraries. Genom ic DNA from
Synechococcus PCC 7942 was digested to completion with BamHl and Hindlll and electrophoresced on a 0.8% agarose gel. The region of gel containing restriction fragments ranging from 1.9-2.5 kb was excised and the DNA gel purified using the
QlAEX II gel extraction system. Plasmid pUCI9 (Gibco-BRL) was digested with the same two restriction enzymes, electrophoresced and gel purified in a simitar fashion.
Restriction enzyme digested DNA fragments from Synechococcus PCC 7942 and pUC19 were used in a ligation reaction at a 2:l molar ratio, respectively, as described by Sambrook et al. (1989). Restricted DNA was incubated with 0.1 Weiss units of T,
DNA Ligase (New England Biolabs) for 16 hours at 16°C. E. coli DH5a competent cells were transformed with a volume of the ligation reaction as per the instructions of the supplier. Cells were plated ont0 LB plates containing 100 pg ml" ampicillin and 50 pg ml" X-gai, and incubated at 37OC overnight. A Synechococcus PCC 7942 BamHI-Clal subgenomic library was constructed in a similar fashion, using a gel slice containing restriction fragments from 4.5-5.5 kb, and pBluescript II WS (Stratagene) as the vector.
Transformants were screened directly using colony lifts and colony hybridization, as described by Sambrook et al. (1989). Nitrocellulose membranes (Schleicher and
Schuell) were placed ont0 transformation plates, and transfer of cells to the membrane was carried out for approximately 2 minutes. The membranes were placed ont0 fresh
LB-ampicillin plates, and incubated for 12 hours at 37°C. Membranes were then treated with 10% SDS for 3 minutes, a denaturing solution (0.5% NaOH, 1.5 M NaCI) for 5 minutes, a neutralizing solution (1.5 M NaCI, 0.5 M Tris-HCI [pH 7.41) for 5 minutes, and 2 X SSC for 5 minutes. Treated membranes were allowed to air-dry for 30 minutes before being vacuum dried for 2 hours at 80°C. Dried membranes were wetted with 2
X SSC for 5 minutes, and prewashed for 30 minutes at 50°C with 5 X SSC, 0.5% SDS, and 1 mM EDTA (pH 8.0). Cellular debris was removed by gently wiping the surface of membranes with Kimwipes, and washing in 2 X SSC. Membranes were prehybridized and hybridized as described previously. Positive clones were identified based on the strength of the radioactive signal obtained after autoradiography. A secondary screen was petformed on the clones identified from colony hybridizations. Groups of clones were used in rapid cell disruptions, separated using agarose gel electrophoresis, and
screened using Southern hybridization. A tertiary screen was performed to narrow
down the number of positive clones further. This time individual clones were resolved
on agarose gels separately and screened using Southern hybridization. This
methodology was used to identify both pBH22 and pBC5O.
Construction and Screening of Lambda Phage Library. A sub-genomic Lambda
phage library of Synechococcus PCC 7942 was constructed and screened using the
Lambda ZAPll (Stratagene) system as per the manufacturers instructions. Lambda ZAP
II arms, supplied pre-digested with EcoRl and dephosphorylated with calf intestinal
phosphatase, were ligated to 150 ng of EcoRl digested genomic DNA from
Synechococcus PCC 7942 using 2 Weiss units of T, DNA Ligase. Restriction fragments of EcoRI-digested genomic DNA, ranging in size from 2-1 0 kb, were purified from an agarose gel as described previously. Ligation was carried out for 2 days at 4°C
prior to packaging of phage using the Gigapack III Gold Packaging Extract (Stratagene)
as per the instructions of the manufacturer.
The packaged phage library was screened by first infecting 600 plof OD,,,= 0.5
XLI-Blue MRF' cells with 1.O MLof a 1:2 dilution of the packaged phage library stock.
Infection was carried out at 37°C for 15 minutes. The host cell-phage suspension was
plated ont0 NZY plates (Sambrook et al, 1989) after 7.0 mL of top agar (LB media with
0.7% agarose) was added, and the mixture vortexed briefly. The plates were allowed to cool to allow the top agar to solidify, and then transferred to 37°C for 8 hours. The same procedure was repeated a total of 10 times to obtain 10 plates that would be screened by way of plaque lifts and hybridization.
Nitrocellulose membranes were layered ont0 the surface of plates, and transfer was allowed to proceed for 2 minutes. Asymmetrical orientation holes were made on each plate to facilitate the identification of plaques after hybridization. Subsequent to transfer, membranes were treated by first submerging them in a denaturing solution
(1.5 M NaCI, 0.5 M NaOH) for 2 minutes, and transferring them to a neutralizing solution (1.5 M NaCI, 0.5 M Tris-HCI [pH 8.01) for 5 minutes. The treated membranes were finally rinsed (0.2 M Tris-HCI [pH 7.51,2 X SSC) for 30 seconds before being blotted briefly, and vacuum dried for 2 hours at 80°C.
Nitrocellulose membranes were prehybridized (2 X Pipes buffer, 50% deionized formamide, 0.5% SDS, 100 pg ml" salmon sperm DNA) for 4 hours at 4Z°C in plastic
Seal-A-Meal bags. After the prehybridization solution was removed, hybridization solution (same as prehybridization solution) was added, as was the radiolabelled probe
(prepared as described previously). The nitrocellulose membranes were allowed to hybridize to the probe for 16 hours. The membranes were washed in the same manner as described for Southern hybridizations. Membranes were prepared for autoradiography as described previously.
Positive hybridization signals were correlated with plaques using the orientation marks made previously. Positive plaques were cored and placed in 500 pL of SM buffer
(0.01 M NaCI, 0.0081 M MgS0,-7H,O, 0.05 M Tris-HCI [pH 7.41, 0.01% gelatin) with
20 pL of chloroforrn. The ExAssist/SOLR system, that accompanies the Lambda ZAP --
II vector, was used to excise the pBluescript SK(-) phagemid from the Lambda ZAP II vector, and in turn allow for recovery of the cloned insert of interest. In 50 mL conical tubes, 200 pL of ODm=l .O XL-1 Blue MRF'cells, 100 MLof the cored phage stock, and
1 PLof ExAssist helper phage were combined, and incubated ai 37°C for 15 minutes.
To this mixture was added 3 mL of 2 X M media (0.1 il M NaCI, 10 g L" yeast extract,
16 g Lm'bactotryptone). The resulting mixture was incubated at 37°C for 2 hours with shaking. The tubes were heated at 70°C for 20 minutes, and centrifuged at 2500 rpm for 15 minutes. The resulting supernatant, which contains the plasmid packaged as a filamentous phage particle, was transferred to a sterile tube for storage at 4°C. To plate the rescued phagemid 1pL of the above mentioned supernatant was used to infect 200 pL of OD,,, = 1.O SOLR cells, which were in turn incubated at 37°C for 15 minutes, before being plated ont0 LB ampicillin (50 pg ml*') plates and incubated at 37°C for
14 hours. A few colonies from each plate were picked and the plasmid sizes determined using rapid cell disruptions. Those colonies that contained plasmids in the expected size range were streaked out and used for plasrnid DNA preparations and restriction analysis. Southern hybridization was used to identify the clone of interest.
Insertional Inactivations
A targeted mutant of Synechococcus PCC 7942 was constructed using an antibiotic resistance cassette to disrupt the coding sequence of ORF146. This was achieved by first subcloning an Xbal-Hinalll restriction fragment from pBH22 and ligating it into pUC19, forming plasmid pXH9. Plasmids pXH9 and pHP45R (Prentki and Kirsch, 1984; kindly provided by Dr. John Coleman, University of Toronto) were digested to completion with 6gAl and BamHI, respectively, and electrophoresced .
Plasmid pHP45D contains an cassette (fi-Spcr), made up of the aadA gene
(conferring resistance to spectinomycin) flanked by transcription-translation termination sequences and restriction enzyme sites. Next, the 3589 bp restriction fragment corresponding to linearized pXH9, and the -2.0 kb restriction fragment, corresponding to the excised n-Spcr, were gel purified and ligated, yielding plasmid pXH::ORF146.
Transformation of E. coli DH5a yielded spectinomycin resistant colonies, that were picked, streaked out, and used for plasmid DNA preparations.
Cells of Synechococcus PCC 7942 were transformed with pXH::ORF146 using a method described by Laudenbach and Grossman (1991). Standing culture cells of
Synechococcus PCC 7942 were concentrated to approximately 1 X IOg celis ml'' before adding pXH::ORF146 to a final concentration of 1 pg ml-'. Transformation was initiated by incubating the cells in the dark for 5 hours at 30°C with agitation, before transferring them to the light (30 pmol (photons) m'20s'' (PPFD)) for 16 hours at 30°C with agitation. Standard BG-Il-agar media, supplemented with 50 pg ml"
Spectinomycin was used to plate out 100 pL aliquots of the transformation reaction.
Plates were incubated at 30°C in Seal-a-Meal bags, supplied with 5% CO, (v/v), under constant illumination at 50 pmol (photons) m-26' (PPFD). Transformants were plated out three times, to ensure complete segregation of the intended lesion before being analyzed further. A similar strategy was used to construct an insertionally inactivated - - mutant of ORF440. Plasmid pP12 was constructed by subcloning an 1197 bp Psf fragment of clone pE56 into pUC19. Both pP12 and pBSL128 (Phabagen) (Alexeyev et a/.,1995) were digested to completion with Clal and electrophoresced. Plasmid pBSL128 contains an C2 cassette made up of the nptll gene, conferring resistance to kanamycin, flanked by transcription-translation termination sequences and multiple cloning sites. The fi-Knr cassette from pBSL128, and pP12 were both isolated by gel purification, as described previously. After ligation and transformation, kanamycin resistant colonies were picked, streaked out, and used for plasmid DNA preparations.
Cells of Synechococcus PCC 7942 were transformed with pPI2::ORFUO as described above, except that transformation plates were supplemented with 10 pg ml'' kanamycin. To verify the genotype of the insertionally inactivated CS1 and CA1 mutants, genomic DNA was isolated and used for Southern hybridization analysis as described previously.
RNA Gels and Northern Analysis
Agarose gel electrophoresis of isolated RNA from Synechococcus PCC 7942 was performed essentially as outlined by Sambrook et al. (1989). Gels were prepared by dissolving an appropriate amount of agarose to obtain a final gel concentration of
1.2% [w/v]. After allowing the mixture to cool to 60°C,formaldehyde and 5 X gel running buffer (FGRB) (0.1 M MOPS, 40 mM sodium acetate, 5 mM EDTA [pH 8.01) were added to 2.2 M and 1 X, respectively. After rnixing, the gel was cast and allowed VL to set for a minimum of 30 minutes.
RNA samples were prepared for electrophoresis essentially as described by
Sambrook et al. (1989). RNA was added to RNase-free microfuge tubes already containing 2.0 PL of 5 x FGRB, 3.5 pL of formaldehyde, and 10 MLof formamide. The final volume of the mixture was brought up to 20 pL with DEPC-treated H,O. Samples were incubated at 70°C for 20 minutes, chilled on ice, and centrifuged briefly. Ethidium bromide (1 pL of a 1 mg ml-') was added to samples only if they were to be visualized.
To each sample was also added 2 pL of a DEPC-treated formaldehyde gel-loading buffer (50% glycerol, ImM EDTA [pH 8.01, 0.25% bromophenol blue, 0.25% xylene cyan01 FF). Before loading samples, the gel was run for 10 minutes at 70 V, and the wells flushed with the running buffer. Samples were loaded and electrophoresis carried out for 2 hours at 60 V in 1 x FGRB. Lanes containing samples that were stained with ethidium bromide were cut out and visualized using the Gel-Docl000 lmaging system.
The amount of RNA loaded in each lane was estimated using spectrophotometry and visual inspection of ethidium bromide stained gels. An RNA ladder was also used to estimate the size of RNA transcripts after hybridization and autoradiography.
Formaldehyde gels were prepared for Northern blotting by repeated rinsing with
DEPC-treated H,O. Gels were blotted to Zeta-Probe (Bio-Rad) nylon membranes according to the manufacturers instructions. The method used was essentially the same as that used for Southern blotting except that acid depurination, alkaline denaturation, and neutralization were not employed. Capillary transfer of RNA to the membrane was carried out for 14-18 hours, followed by rinsing the membrane in 2X SSC. Efficiency of transfer was determined by staining gels with ethidium bromide for
45 minutes and then visualizing the gel under UV light. RNA was fixed to the membranes by vacuum drying at 80°C for 30 minutes. Northern hybridization of radiolabelled DNA probes to blots was performed as described for Southern hybridizations.
Measurement of Photosynthetic O, Evolution
Photosynthetic O, evolution was measured using a therrnostatted Clarke-type electrode (Hansatech, Norfolk, U.K.). Cells were washed three times by centrifugation
(Beckman Microfuge E) with low-C, buffer (Espie and Canvin, 1987). After the final wash, the cell pellet was resuspended in 1.5 mL of the same buffer, and transferred to the 30°C thermostatted chamber of the 0, electrode. The cell suspension was gassed with N, to remove O, and constantly stirred with a magnetic flea. After gassing with N, the chamber was closed to prevent contamination of the cell suspension by atmospheric gases. Additions to the cell suspension were made through a capillary bore in the plug, using Hamilton syringes. Actinic light to drive photosynthesis was provided at 100 pmol (photons) m'2 s" (PPFD). After cells had reached the CO, compensation point, photosynthesis was initiated by introducing Ci (in the form of
KHCOJ or KOCN to the cell suspension. After an experiment was completed, the cell suspension was collected and [Chl a] determined spectrophotometrically at 665 nrn after extraction in methanol as described by MacKinney (1941). Fluorometry
Chlorophyll a fluorescence quenching of cell suspensions was measured using a pulse amplitude modulated fluorometer (H-Walz, Germany, PAM 101, Schreiber et a/., 1986) as described by Miller et al. (1991 ). Cells were washed and resuspended in
25 mM BTP/HCI (pH 8.0) buffer, and placed in a 30°C thermostatted Clarke-type electrode. This facilitates the sim ultaneous measurement of photosynthetic O, evolution and fluorescence quenching. The cell suspension was continuously stirred with a magneticflea. A weak pulse modulated red light beam (5 pmol (photons) mq s" at 100 KHz) was used to monitor fluorescence yield. The light was delivered to the cell suspension using a four-armed fiber optic cable. A photodiode set to detect wavelengths in the range of 685 nm (chlorophyll a fluorescence) was used to detect fluorescence. Actinic light to drive photosynthesis was supplied at 100 pmol (photons) me*s'' (PPFD). Minimal fluorescence yield (F,) was determined by illuminating dark adapted cells with the weak pulse modulated light beam. Maximum fluorescence yield
(Fm*)was determined by delivering a saturating light flash (1600 pmol m'* s'' for 1 s) to illuminated cell suspensions at the CO, compensation point. Changes in fluorescence are expressed as a percent maximum of the variable fluorescence (Fv*), where Fv* = Fm*-F,. Fluorescence yields and initial rates of fluorescence quenching have been correlated with the size of the interna1 Ci pool and initial rates of Ci pool formation (Miller and Canvin, 1987; Miller et al., 1988; Miller et a/, 1991; Espie et al.,
1991; Crotty et al., 1994). In the context of the current work, chlorophyll a fluorescence - - quenching was used to determine if a OCNxJependent interna1 Ci pool was formed upon the addition of KOCN to Ci depleted cells.
Mass Spectrometry
Dissolved l2CO2in cell suspensions was measured using a VG gas analysis magnetic sector mass spectrometer (Middlewich, England) connected to a membrane inlet system as described by Miller et al. (1988). Cells suspensions (6 mL), washed three times with BTPIHCI (pH 8.0) buffer, were incubated in a 30°C thermostatted glass cuvette and continuously stirred using a magnetic flea. To avoid contamination of the cell suspension by atmospheric gases the suspension was purged with N, until a
Plexiglas plug was used to seal off the cuvette. Additions of KHCO, and KOCN were made with Hamilton syringes through a capillary bore in the plug. The cell suspension was separated from the vacuum chamber of the mass spectrometer by a thin gas permeable dimethyl silicone rubber membrane supported by a metal grid. It is located at the tip of the gas inlet tube and is inserted into a port on the reaction vessel. The dimethyl silicone inlet membrane is permeable to dissolved CO2 but not HCOj or COP.
Therefore, the mass spectrometer measures only dissolved [CO,] directly, not [HCO;] or [COA. Actinic light to drive photosynthesis was provided at 21 0 pmol (photons) m'2 s'' (PPFD).
The mass spectrometer was calibrated for CO, by the addition of specific amounts of kCO, to pH 8.0 buffer. The equilibrium [CO,] was calculated as described by Buch (1960) taking into account the temperature, pK, pH, and ionic strength of the
buffer. At pH 8.0, 1.56% of the K,CO, will be in the form of CO, and with the addition
of 25 mM NaCl 1.39% of &CO, will be CO,. The rnass spectrometer was also used to
determine the extent of CO, contamination of KOCN stock solutions, since OCN- can
spontaneously breakdown to form CO, and NH,. Specific amounts of KOCN were
added to the same buffer and the mass spectrometer used to determine the amount of
CO, in the stock solution of KOCN. At a final dilution of 1 mM, the KOCN solution was found to contain -5 pM Ci.
Cyanate Transport Assays
Transport and intracellular accumulation of OCN' was assayed using the silicone fluid centrifugation technique with K0I4CN as substrate (Miller et aL, 1988). Standing culture cells of Synechococcus PCC 7942 (wild type, CS1 and CA1) were harvested when they had reached a concentration of 5-7 pg ch1 a ml", and concentrated to approximately 15 pg ch1 a ml". The concentrated cell suspension was washed 3 times with BTP/ HCI pH 8.0 buffer, and placed in 30°C- thermostatted Clarke-type O, electrode as described previously. Light was provided at 100 pmol (photons) m2 s"
(PPFD). After the addition of 25 mM Na', the cell suspension was allowed to fix available Ci until net 0, evolution ceased. Once the compensation point had been
reached, KO14CN (SA= 2.5 to 5.0 pCurries pmol") was added to a final concentration of 2 mM. After 5 and 10 minutes, 3 x 100 ML samples were removed from the cell suspension under constant illumination and layered on top of (frorn top to bottorn) 100
PLsilicone fluid (1.75 AR20 : 1 AR200; Wacker Chemie, Munich, Gerrnany) and 100 pL killing solution (2 M KOH in 10% methanol) contained in a 400 pL microcentrifuge tube. Cells were centrifuged through the silicone fluid and into the killing solution in a
Beckman rnicrocentrifuge B at 12 000 x g for 1 minute. Tubes were subsequently frozen and stored in liquid nitrogen until al1 samples were ready to be processed.
Processing involved cutting the frozen tube at the silicone oil-killing solution interface.
The bottom of the tube containing the killing solution and the cell pellet was kept for further processing. Once thawed, the cell pellet was resuspended in the killing solution and transferred to a new tube. The cut tube was washed once with 100 pL of basic water, which was then added to the ce11 suspension. Two 50 pL samples of the ce11 suspension were transferred to two separate tubes. To one of the tubes, 1.2 mL of
Universol (ICN) scintillation cocktail was added and to the other 100 pL of 4 M acetic acid. The former was placed in a liquid scintillation counter (Wallac 1400) and its radioactivity measured (total cell pellet-associated I4Ccounts), whereas the latter was baked overnight at 80°C (acid stable counts). After baking, the remaining pellet was resuspended in 100 HL of water, and radioactivity counted after the addition of scintillation cocktail. The difference between the total cell pellet counts and acid -stable counts, corrected for extracellular contamination, represents the interna1 pool of unreacted 14C. RESULTS
Construction of Subgenomic Libraries and Isolation of a Genomic Region
The nucleotide sequence of cynS from E. coli KI2 was used to construct oligonucleotide primers (20- mers) for polymerase chain reaction (PCR). The sequence of the primers was identical to corresponding regions of cynS, and were designed to arnplify a 483 bp portion of cynS. After PCR and electrophoresis, the expected 483 bp product was obtained, gel purified and blunt-end ligated into pUC18 using the
Sureclone (Pharmacia) system. Subsequent to transformation, plasm id DNA was isolated and used for automated nucleotide sequencing using Ml3 fotward and reverse primers. The sequence that was obtained was 100% identical to that of cynSfrom E. coQGenBank Accession Nurnber Ml7891). This insert would serve as the probe for preliminary Southern hybridization analysis of Synechococcus PCC 7942 genomic
DNA.
Genomic DNA extmcted from Synechococcus PCC 7942 was digested with restriction endonucleases, electrophoresced, Southern blotted and hybridized with the cloned PCR product of E. coli cynS. Analysis of the autoradiogram (Figure 1A) indicated that the probe hybridized to restriction fragments in each lane, ranging in size from approximately 2.2 to 22 kb in length. It should also be noted that in each lane only one band was apparent. An approximately 2.2 kb band was obtained from the digestion of Synechococcus PCC 7942 genomic DNA with BamHl and Hindlll. The restriction 39
Figure 1. Southern hybridization analysis of genomic DNA from Synechococcus PCC
7942. lsolated genomic DNA was digested with appropriate restriction enzymes, electrophoresced on 0.8% agarose gels at 60 V for 2 hours, and Southern blotted.
Hybridization and post-hybridization washes of the blots were performed at 65°C. A)
Restriction enzyme digested genomic DNA from wild type Synechococcus PCC 7942 cells probed with a 483 bp region of cynS from E. coli K12. B) Restriction enzyme digested genomic DNA from wild type cells probed with the 1287 bp BamHI-Xbal region of pBH22. The positions of molecular weight standards are indicated to the left of A) and B). Restriction enzymes used are indicated (8, BamHI; Bg, Bg111; C, Clal; El EcoRI;
H, HinuIll; K, Kpnl; P, Psll; X, Xbal). BBgC H K Pm
E H BBgXP fragments obtained from BamHI-Hindll ranging in size from 1.9 to 2.5 kb were isolated for construction of a pUCl9 subgenomic plasmid library for the identification of a cynS homolog in Synechococcus PCC 7942. Colony lifts and hybridization were used to screen E. coli DH5a cells transformed with the Synechococcus PCC 7942 pK19 subgenomic library. A single clone, designated pBH22, was isolated and plasmid DNA was obtained for sequencing. Both strands of pBH22 were sequenced without arnbiguities. Initial nucleotide sequence obtained from the Hindlll end of the pBH22 insert, using Ml3 reverse sequencing primers was found to be similar to E. coli cynS
(discussed later). Initial sequence obtained from the BamHl end of the insert was similar to genes that are members of ABC transport operons. To extend the genomic region already isolated, two other subgenomic libraries were screened.
Sequences upstream and downstream of the region isolated from pBH22 were obtained by screening two separate subgenomic libraries. To identify the region upstream of BamHI, Southern analysis was performed, using the BamHI-Xbal region of pBH22 as a probe (Figure 2). Southern analysis (Figure 1B) revealed single bands in each of the lanes. The bands ranged in size from -3.5 kb to 11 kb. The lane containing genomic DNA digested with EcoRl gave a band of approximately 5.6 kb. An available Lambda Zap II EcoRl subgenomic library was used to isolate clone pE56.
Restriction analysis of the recovered phagemid clone indicated that it contained an insert of -5.6 kb. Isolated plasrnid DNA was used for automated sequencing and initial nucleotide sequence data obtained with Ml3 universal primers was identical to that of the overlapping region from pBH22. Figure 2. Organization of the genomic region isolated from Synechococcus PCC 7942.
A) Genomic clones from Synechococcus PCC 7942 and their relative position with
respect to the cyn operon. B) Shaded boxes indicate the identified open reading frames. The bold arrow (below) indicates the respective position of the open reading frames, the direction of transcription and the gene designations. Probes used in this
study are indicated by horizontal lines above the region that they correspond with: a) the 687 bp Psfl-Clal region of pP12 used for Southern analysis of CAI; b) the 1197 bp
Psti region of pP12 used for Northern analysis; c) the 1287 bp BamHI-Hindi11 region
of pBH22 used for Southern analysis; d) the 187 bp EcoRI-BgAl region of pBH22 used for Southern analysis of CSI. e) the 465 bp EcoRI-Hindlll region of pBH22 used for
Northern analysis. Restriction enzyme site positions are indicated (B, BamHI; Bb, Bbyl;
Bg, BI;C, Clal; E, EcoRI; H, Hindll; P, Psd; X, Xbal). Sites where antibiotic
resistance cassettes were inserted, the type of cassette used and corresponding
mutant strain generated are indicated by filled arrow heads (A). E Bg HEH
1 1 Knr (CAI) Spc' (CS) 245 bp
ORF 440 ORF289 ORF146 44
The region downstream of Hindlll was identified by constructing and screening a subgenomic pBluescript II WS BamHI-Clal library of Synechococcus PCC 7942.
Southern analysis of Synechococcus PCC 7942 genornic DNA with the cloned PCR product of cynS from E. coli indicated that a single hybridization signal at -5 kb was obtained with genomic DNA digested with BamHl and Clal (results not shown). The region from 4.5 to 5.5 kb was isolated for construction of a subgenomic plasmid library.
Colony lifts and hybridization were used to identify clone pBC50. Restriction analysis of pBC50 indicated that it contained an insert of - 5.0 kb and initial sequence was identical to that which was obtained for the overlapping region of pBH22.
Sequence Analysis of pBH22, pE56 and pBC50
A restriction map (Figure 2) of the genomic region cloned from Synechococcus
PCC 7942 was constructed using the nucleotide sequence data obtained from pBH22, pE56 and pBC5O (Figure 3). Initial sequence data was analyzed using the BLASTN
(Basic Local Alignment search 1001for nucleotide sequences), and BLASTX programs
(Altschul et aL, IWO). Results from BLASTN analysis of pBH22 indicated three reg ions that exhibited significant sequence similarity to non-redundant sequences at GenBank,
EMBL, DDBJ, and PDB. Results from BLASTX also indicated three regions with significant sequence similarity to GenBank CDS translations, and peptide sequences at PDB, Swiss-Prot, and PIR. BLASTX operates by translating a nucleotide sequence in al1 six reading frames and comparing the resulting protein sequences to existing Figure 3. Nucleotide and deduced amino acid sequence of the 4453 bp region identified in Synechococcus PCC 7942. Arrows (>) indicate the direction of transcription of the identified open reading frames. Amino acid residues conforming to the ATPIGTP-binding site A motif are indicated in bold type. Amino acid residues conforming to the ABC transporters family signature are underlined. Putative ribosome binding sites are indicated by a line over the sequence. Putative hair-pin structure sequences are indicated by a double line under the sequences that are inverted complements of each other. CGTGTCGCTGTCAGGATGCTATTCGTCGACGACCGATCACGATCGCTTGGGAGATATGCCGCGACCTGAGATCTTGATGGCAGATTTTCGATGG 100 GTTTATTCAAACCTGAATCGATTGCGATCGCCGATCGTCTGACGACTTCAGAGTTGCGCCGCCTTGTTCTTTTGCTTCCCGGGGATCCTGA 200
ACAGGGAAGC~ATGCTGAGAGCTTTGGCTGCAGAAGCAGGTGGTTTAGAT~AGG~ATTTG~G~TG~TTTTGGA~~G~G~GGGT~GTTTTTT~AG~~300 MLRALAAEAGGLDQAFAAAFGPQAGRFFSN 0-440 > CATTCAAGCTAGTGGCGGGGTCAGTCGTCGGGTTTTCCTGCGCCTTGGTCGTTGGTGCAGCTCTCGTGACGCTGACCCTGTGCCCCAGGCTC 400 IQASGGVSRRVFLRNLVVGAALVTLTNCAQQAQ
CAACCCGATAGCCCAACCACGACTGGCAGTGGCAATTTTAGUCGGATCTGmGGTTGGCTTTATCCCGATTACCTGCGCCACTCCGATCATCATGT 500 QPDSPTTTGSGNLEKTDLKVGFIPITCATPIIM
CAGATCCCTTGGGCTTCTATCAGAAATATGGCTTGAAAGTCCAAGTTGTGAAGATGCCGAGCTGGGGGGCAGTTCGAGATTCTGCGATCGCAGGCGAATT 600 SDPLGFYQKYGLKVQVVKMPSWGAVRDSAIAGEL
GGATGCCTATCACATGCTGGCACCGATGCCGATCGCGATGACCTTGGGTCTTGGCTCAGCTCCCTTCAGTGTCAATGTTAGCCAGTATTGTATTC 700 DAYHMLAPMPIAMTLGLGSAPFSVKLASIENIN
GGTCAGGCGATTACGGTTGCCAAACGTCACCTTGGCCTGC800 GQAITVAKRHLGKVKEAKDFKGFVIGVPFPFSM
ATAACCTGCTGTTGCGCTACTATCTCGCTGCTGGTGGTTTGTCCCGATACCGATGTCCAAATTCGGCCAGTTCCCCCGCCAGATAGTATTGCTCAGCT 900 HNLLLRYYLAAGGLNPDTDVQIRPVPPPDSIAQL
CGTCGCAGGTGATATCGATGCGATGCTGATGCCCGATCCCTTTAATCAGCGGGCAGTGTATGGATGCTGGCTTTATTCATCTGTTCTGTT 1000 VAGDIDAMLMPDPFNQRAVYEDAGFIHLLTKEI
TGGAATGGTCATCCTTGCTGTGCATTTGCAGCAGGTGAGCCTTGGATTCGTCCCTACGTTCCGAGCGCTTCGCTTATTGAAGC 1100 WNGHPCCAFAAGEPWIQENPNTFRALNKAIIEA
CTGGTTATGCCAGTAAGGCCGAAAATCGTGCTGAGATTGCCmGGCTATTTCTAGCCGTCAGTACTT~TCAACCACCCGMGTCGTGGMGCTGTGCT1200 TGYASKAENRAEIAKAISSRQYLNQPPEVVEAVL
GACCGGTAAGTTCCCCAATGGTCMGGTCAAGAACTGGATGTTCCCGATCGCATTGACTTCTCCCTACCCATGGCAGAGCTTTGCCCTGGATTC 1300 TGKFPNGQGQELDVPDRIDFNPYPWQSFANWIQ
TCGCAGCTAGTGCGTTGGGATCTGGGTAAAGCTGCCGGTGTGATCCAGCCCGATCAGTACGACGCGGTCAGGCTTTACCTGACGACTGAAGCAC 1400 SQLVRWDLGKAAGVIQPDQYDKNGQAIYLTTEA
AAACCCTCGAGAAGGAAGTGGGCCTGCAGCCGCCGACTGATCGCTTA 1500 QTLEKEVGLQPPTEIYREEKLAYDTFNPQDPVAY CCTCGCATCTCAAAAGCAGAAATACGGGAGATAAACACAACTTATGGTGAGAACTCCTGTACCGCTTTACCTACGTTGGGCGGTCTCCATCCTCAG~GTG16O O LASQKQKYGR* MVRTPVPLYLRWAVSILSV ORF263 > CTTGCGTTCCTAGCCATTTGGCAAATTGCGGCAGCTTCAGGATTTTTAGGCCTTTTCCTGGCTCCCTGCGCACTTTGCAGGATTTGTTTGGATGGC 1700 LAFLAIWQIAAASGFLGKTFPGSLRTLQDLFGW
TTTCAGATCCCTTCTTTGATAACGGCCCCAATGACTTCC1800 LSDPFFDNGPNDLGIGWNLLISLRRVAIGYLLAT
AGTTGTTGCAATTCCTTTGGGGATTGCAATCGGTATGTCGGCGCTAGCTTCCAGTATTTTTTCGCCCTTTGTGCAACTCCTGAAGCCAGTTTCACCTTTG 1900 VVAIPLGIAIGMSALASSIFSPFVQLLKPVSPL
GCCTGGTTGCCGATTGGTCTCTTCTTATTCCGAGATTCGGTTGACGGGTGTTTTTGTCATCCTGATTTCGAGTCTGTGGCCAACGTTGATCCACAG 2000 AWLPIGLFLFRDSELTGVFVILISSLWPTLINT
CGTTTGGGGTGGCGMTGTCAATCCTGACTTTTTGMGGTTTCGCAATCTTTGGGAGCTAGTCGTTGGCGCACGATTCTGAAGGTGATTCTGCCCGCAGC 2100 AFGVANVNPDFLKVSQSLGASRWRTILKVILPAA
ATTGCCCAGCATCATCGCGGGAATGCGGATCAGCATGGGCATTGCTTGGCTGGTCATTGTGGCAGCAGAGATGCTGTTGGGUCAGGmTTGGCTATTTC 2200 LPSIIAGMRISMGIAWLVIVAAEMLLGTGIGYF
ATTTGGAATGAGTGGAATAACCTATCACTTCCTAATATTTTCTCGGCCATCATCATCATTGGGATTGTTGGCATTCTTCTCGACCAAGGCTTCCGTTTTC 2300 IWNEWNNLSLPNIFSAIIIIGIVGILLDQGFRF
TTGAGAACCAGTTTTCTTACGCAGGCAACCGATCCCATGATTTCTGAAGCTGTGCCAGCCAAGGAGGAGACAGGGCAGGCTCTTGCTGATTGAGCA 2400 LENQFSYAGNR* MISEAVPAKEETGQAQLLIEQ ORFS 8% AGTTGGCAAAGTTTTTACTGTCAATTCACCTTCACCTTCTCTCCTCGATCGCCTTCGACAGCGATCGCCCAAACGCTACGTTGCATTAGAAGATGTCAACCTCACG2500 VGKVFTVNSPSLLDRLRQRSPKRYVALEDVNLT
ATCGCGTCGAACACATTTGTCTCGATTATTGGCCCTTCGGGTTGTGGTTCCCCTTCTCCTTGATTGCTGGCCTTGATTTACCCGTCTGGCC 2600 IASNTFVSIIGPSGCGRSTLLNLIAGLDLPTSG
AGATTCTGCTGGATGGTCAATTCACCCGCATTCGATCGCCGGGGCCCGATCGTGGCATCGTCTTCCAGAACTATGCCCTGATGCCCTGGATGACCGCGCTTGAGAA2700 QILLDGQRIRSPGPDRGIVFQNYALMPWMTALEN
TGTCATCTTTGCAGTTGAAACGGCGCGCCCAAACCTGAGCTCCCGCTCGCGGTGGCACGAGAGCATCTAGAGCTGGTGGGTTTCCGCT 2800 VIFAVETARPNLSKSQAREVAREHLELVGLTKA
GCCGATCGCTATCCGGGCCAAATTTTCAGGGGGGATGmCAGCGCGTAGCGATCGCCCGTGCCCTCTCCATCCGTCCTmGCTCCTGCTGATGGATGMC 2900 ADRYPGQISGGMKQRVAIARALSIRPKLLLMDE O O O m wv) 8 8 E-t Uk Y3 l3 l4 Uw E > w i? > "wy= ES. cl izU U fx a 8 riz Ec3" wçr; wU E J U E 3 3 il w w3 a 3U U " LI 3U U ww E Uçr; u Y"UVI U" U a U w " x 3u S B u" Pi U C3 > "w w E LI E" UPi sequences. Sirnilar analysis was performed on sequence data obtained frorn pE56. The results obtained from BLASTN and BLASTX were used to identify four open reading frames (ORF's) that are closely spaced in relation to each other. The four ORF's are presented as ORFI 46,ORF289,ORF263, and ORF440 (Figure 2) and also designated as cynS, cynD, cynB, and cynA, respectively.
ORF146. A putative ORF coding for a protein of 146 amino acids was first identified based on its nucleotide sequence similarity to cynSfrom E. coli. The sarne
ORF also had significant sequence similarity to a potential cynS homolog from the cyanobacterium Synechocystis PCC 6803, in which the entire genome has been recently sequenced (Kaneko et al., 1995; Kotani et al., 1994; Kotani et al., 1995). As shown in Figure 2, ORF146 is located at the HinoIll end of the insert of pBH22, downstrearn of the other three identified ORF's. The identified ORF initiated with an
ATG at position 3219 (Figure 3) and terminated with a TAA at position 3260. Located ten bases upstream of the initiator, a putative ribosome binding site was identified with the sequence GGAGA (Figure 3). Analysis of the upstream region did not identify any obvious promoter sequence characteristic of other cyanobacterial genes or those of E. coli a7*or 82promoter consensus sequences.
The FASTA program (FASTA version 3.0t71 November 1996) (Pearson and
Lipman, 1989) was used to align the deduced amino acid sequence of ORF146 to that of protein sequences at Swiss-Prot (Figure 4A). FASTA analysis indicated that
ORFI 46 aligned with, and exhibited 43% sequence identity to CynS from E.coli in a ORF14 6 MTSAITEQLLKAKKAKGITFTELEQLLGRDEVWIASVFYRQSTASPEEAEK 51' ...... I :I 1:::I:I::: 1 :I::::::: [ : ::::I CynS MIQSQINRNIRLDLADAILLSKAKKDLSFAEIADGTGLA~FWULLGQQALPADMRL 60"
ORF146 LLTALGLDLALADELTTPPVKGCLEPVIPTDPLIYRFYEIMQGLPLKDVIQEKFGDGI 111' : : [:II : 1 I::II:: 11111 :11111::1111 :l1::::1111111 CynS VGAKLDLDEDSILLLQMIPLRGCIDDRIPTDPTMYRFYEMLQVYGTTLKALVHEKFGDGI 120
ORF146 MSAIDFTLDVDKVEDPK-GDRVKVTMCGKFLAYKKW 146' :[[~:~:~~~:~~:[~:[:l: :l: l[:l: 1 CynS ISAINFKLDVKKVADPEGGERAVITLDGKYLPTKPF 156"
ORF289 MISEAVPAKEETGQAQLL-IEQVGKVFTVNSPSLLDRLRQRSPKRALEDLTAST 59' ... . :I: 1 :: : :: : :I 1:I I[I:IIIII:: :: NrtD MTAILPSTAATVNTGFLHFDCVGKTFP--TP------RGP--YVAIEDVNLSVQQGE 47 ' ' P-loop ORF289 FVSIIGPSGCGKSTLLNLIAGLDLPTSGQILLDGQRIRSPGPDRGIWQWALMPWTAL 119' 1: :Il 11111111111::l:: 1111 : 1111 1: 11111 :lIIII:I:Il :1 NrtD FICVIGHSGCGKSTLLNLVSGFSQPTSGGVYLDGQPIQEPGPDRMWFQWSLLPWKSAR 107" ABC signature ORF289 ENVIFAVETARPNLSKSQARWAREHLELVGLTKAADRYPGQISGGMQRVAIARALSIR 179' 4: :11::111:11 1: ICI: :IIIIIIII:I : 1 1:11111111111111111 NrtD DNIALAVKAARPHLSTSEQRQVVDHHLELVGLTEAQHKRPDQLSGG~QRVAIARALSIR167-
ORF289 PKLLLMDEPLVPWMPSTRGYLQEEVLRIWEANKLSVVLITHSIDLLLSDRSRGP 239' ~::~::~~~:1: lllI:I III : ~l~~lll~lllll~l~ll~l~l~Il NrtD PEVLILDEPFGALDAITKEELQEELLNIWEEARPTVLMITHDIDEALFLADRVVMMTNGP 227''
ORF289 RATIREVIDLPAVRPRQRSVIEEDERFVKIKLRLEEHLFNETRAVEEASV 289' 111 11:::l 11 1:1 :: [I 1::::: : : 1: NrtD AATIGEVLEIPFDRPREREAVVEDPRYAQLRTEALDFLYRRFAHDDD 274"
Figure 4. Alignment of the deduced amino acid sequence of ORFI 46 with that of CynS from E. coli KI2 (A), and ORF289 with that of NrtD from Synechococcus PCC 7942 (B) as optimized by the FASTA program (Pearson and Lipman, 1988). Vertical lines indicate aligned and identical amino acid residues. Colons indicate conserved amino acid replacements. The P-loop motif (ATPIGTP binding motif A) and ABC transporter family signature sequence are also indicated in B), in bold lettering. 142 amino acid overlap. Over the same number of residues ORF146 exhibited an additional 27% sequence conservation with CynS. Over a 144 amino acid overlap,
ORFI 46 exhibited 77.1 % sequence identity to a potential CynS-homolog from
Synechocystis which was also identified based on its similarity to CynS from E. coli
(alignment not shown).
ORF289. Nucleotide sequence analysis of the region upstream of ORFI 46 indicated that it shared significant sequence similarity to genes that code for ATP- binding proteins that are members of ATP-binding cassette transporters (ABC transporters). An open reading frame of 289 amino acids was identified, that initiated with an ATG at position 2339 (Figure 3) and terminated with a TAG at position 3209,
10 bases upstream of ORF146. Analysis of the upstream region did not identify putative promoter sequences characteristic of other cyanobacterial genes or those of
E. colifl or CPpromoter consensus sequences.
FASTA analysis of ORF289 indicated that it's deduced amino acid sequence aligned with several ATP-binding proteins that are members of ABC transport systems.
ORF289 aligned with and exhibited 50.0% sequence identity to NrtD from
Synechococcus PCC 7942 over 279 amino acids (Figure 48). A search of the
PROSITE database of protein motifs and patterns (Bairoch, 1992) identified two motifs that are characteristic of proteins that are members of ABC transporters (Figures 3 and
4B). The first was a P-Loop motif (ATPIGTP-binding site motif A, PROSITE accession number PS00017) extending from position 2531 to 2554 (Figure 3), encompassing a total of 8 arnino acids. The second motif that was identified was the ABC transporter family signature (PROSITE accession number PS00211), extending from position 2822 to 2865 (Figure 3), including a total of 15 amino acids. These two motifs were also in alignment with the same motifs from NrtD (Figure 4B, bold type), and with the exception of a single residue per stretch, were identical in sequence.
ORF263. Analysis of nucleotide sequence data from the region directly upstream of ORF289 indicated that it shared significant sequence similarity to genes that code for integral membrane proteins that are members of ABC transporters. An open reading frame of 263 arnino acids was identified, terminating 3 bases upstream of the start of ORf289. The deduced amino acid sequence of ORF263 initiated with an ATG at position 1544 (Figure 3) and terminated with a TAA at position 2336. A putative ribosome binding site was identified 10 bases from the start, with the sequence
GGAGA (Figure 3). Analysis of the upstrearn region did not identify putative promoter sequences characteristic of other cyanobacterial genes or those of E. coli~~~or 42 promoter consensus sequences.
FASTA analysis indicated that the deduced amino acid sequence of ORF263 aligned with and shared significant sequence identity to many hydrophobic integral membrane proteins. In particular, it shared 43.4% sequence identity to NrtB from
Synechococcus PCC 7942 over a span of 236 amino acids (Figure 5). Hydropathy analysis of the deduced amino acid sequence using the method of Kyte and Doolittle
(1982) indicated that ORF263 codes for a highly hydrophobic protein (Figure 6). The 1 0RF2 6 3 MVRTPVPLYLRWAVSILSVLAFLAIWQIAAASGFLGKT-FPGSLRTLQDLF 50' 111111: :1 :II: :II : :: : : NrtB LRPPSSVRRSAWVKNPKLKPFLPYVVCLPIFLAIWQVISA--ILGQDRLPGPTW 62" II ORF263 -GWLSDPFFDNGPNDLGIGWNLLISLRRVAIGYLLATWAIPLGIAIGMSALASSIFSPF 109' :: :111111 :: 1:1 ::1111:111111111: ::1 :1 ::Il1 : :: ::I NrtB MPYIVEPFFDNGGTSKGLGLQILISLQRVAIGYLLYGLDPV 122" III ORF263 VQLLKPVSPLAWLPIGLFLFRDSELTGVFVILISSLWPTLINTAFGVPDFLKVSQS 169' :I:I: 1 II1I:II:I::I:I:: :::lll:[:::ll :IlII 1: :: 1: :I:: NrtB IQVLRTVPPLAWPISLMVFQDANTSAIFVIFITAIWPIIINTAVGINQIPDDYNNVARV 182" IV ORF263 LGASRWRTILKVILPAALPSIIAGMRISMGIAWLVIVAAEMLLGTG-IGYFIWNEWN--- 225' 1 1: 11::::I:::l ::11:11::1:111:1111111 : 1 111111: :1 NrtB LKLSKKDYILNILIPSTVP~AGLRIAVGLAWLAIVAAEMLKADGGIGYFIWDAYNAGG 242" v ORF263 NLSLPNIFSAIIIIGIVGILLDQGFRFLENQFSYAGNR 263' : 1 :[: 11: :]:Il: 11: NrtB DGSSSQIILAIFYVGLVGLSLDRLVAWVGRLVSPVSR 277"
Figure 5. Alignment of the deduced amino acid sequence of ORF263 with that of NrtB from Synechococcus PCC 7942 as optimized by the FASTA program (Pearson and
Lipman, 1988). Vertical lines indicate aligned and identical amino acid residues.
Colons indicate conserved amino acid residue replacements. Stretches of sequence with a line above them are putative membrane spanning regions and correspond to the respective hydrophobic regions indicated in Figure 6. Figure 6. Hydropathy profile of the deduced amino acid sequence of ORF263, as determined by the method of Kyte and Doolittle (1982) using a window of 13 amino acid
residues. Roman numerals indicate hydrophobic regions that have been identified as
putative transmembrane reg ions using the TMpred program (Hoffman and Stoffel,
1993). The hydrophobic regions also correspond to the regions identified with horizontal lines in Figure 5.
- - hydropathy profile of ORF263 suggested that it contains five stretches of sequence that are highly hydrophobic in nature (Figure 6). A search of transmembrane protein sequences at TMbase using the TMpred program (Hoffman and Stoffel, 1993) indicated that ORF263 had 5 putative transmembrane helices. Potential transmembrane spanning domains are overlined in Figure 5, and the corresponding regions with large positive hydropathies are indicated in Figure 6.
ORF440.Nucleotide sequence obtained for the region upstream of ORF263, from clone pE56 (Figure 2), was analyzed in the same manner as described previously.
BLASTX analysis indicated that the region contained pockets of sequence that are similar to genes that code for proteins which are periplasmic-binding proteins and/or members of ABC transporters. Further sequence analysis resulted in the identification of a large open reading frame encoding a protein of 440 amino acids (Figures 2 and
3). The identified ORF initiated with an ATG at position 212 and terminated with a TAA at position 1534 (Figure 3) . A putative ribosome binding site was identified 7 bases upstream of the start with the sequence GGAAG (Figure 3). Analysis of the upstream region did not identify putative promoter sequences characteristic of other cyanobacterial genes or those of E. coii~~~or d'*promoter consensus sequences.
The deduced amino acid sequence of ORF440 was analyzed using the FASTA program. ORF440 had sequence similarity and aligned with several periplasmic binding proteins, including NrtA from Synechococcus PCC 7942. The FASTA alignment of
ORF440 with NrtA is presented in Figure 7. The deduced amino acid sequence of ORF440 EAGGLDQAFAAAFGPQAGRFFSNIQASGGVSRRWLRNLWG---AALVTLTNCAQQAQQ 64' [II II I::I II : 1: 1::: :: NrtA MSQFSRRKFL--LTAGGTAAAALWLNACGSNNSS 32"
ORF440 PDS------PTTTGSGNLEKTDLK---VGFIPITCATPIIMSDPLGFYQKYGLK-VQVVK 114' . . 1: :I::: : ::I : : 1:: 1:: ] ::III NrtA TDTTGSTSTPAPSGTSGGDAPEVKGVTLGFIALTDAAPVIIALEKGLFAKYGLPDTKVVK 92"
ORF440 MPSWGAVRDS----AIAGELDAYHMLAPMPIaMTLGL---GSAPFSLASIENINGQAI 167' : 1l:::Il: : 1 1: :::I:I :: 1: : : : I :II:I NrtA QTSWAVTRDNLELGSDRGGIDGAHILSPMPYLLTAGTITKSQKPLPMYILARLNTQGQGI 152"
ORF440 TVAKRHLGKVKEAKDFKGFVIG------VPFPFSMHNLLLRYYLAAGGLNPDT 214' ::::: 1:: : II 1 :I: 1 11 : 1:l :~~:~~~:~::~:: NrtA SLSNEFLAEKVQIKDPKLKAIADQKKASGKLLKAAVTFPGGTHDLWMRYWLAANGIDPNN 212'
ORF440 DVQIRPVPPPDSIAQLVAGDIDAMLMPDPFNQRAVYEDAGFIHLLTKEIWNGHPCCAFAA 274' 1::: ::1:: : 1:: : 1:1 1 : 1: : 1:: 1 1::: NrtA DADLWIPPPQMVANMQTGTMDTFCVGEPWNARLVNKKLGYTAAVTGELWKFHPEKALTI 272"
ORF440 GEPWIQENPNTFRALNKAIIEATGYASKAENRAE1AKAI:SSRQYLNQPPEVVEAVLTGKF 334' 1 ::II:: II 11: II : 1 1: : :: :I:: 1 :: II: NrtA RADWADKNPKATMALLKAVQEAQIWCEDP~LDELCQITAQDKYFKTSWDIKPRLQGDI 332"
ORF440 PNGQGQELDVPD-RIDF----NPYPWQSFANWIQSQLVRWDLGKAAGVIQPDQYDKNGQA 389' I:]: : 1 1: 1 :I::I 1: :: :II 1: : :I ::: NrtA DYGDGRSVKNSDLRMRFWSENASFPYKSHDLWFLTEDIRWGYLPASTDT-LIKRS 391"
ORF440 IYLTTEAQTLEKEVGLQPPTEIYREEKLAYD--TFNPQDPVAYLASQKQKYG 440' 1::: :I : 1: 1 : :1 11:1::1 111 : II NrtA DLWREAAKAIGREQDI--PASDSRGVETFFDGVTFFDGVTFDPENPQAYLDGLKFKAIKA 443"
Figure 7. Alignrnent of the deduced amino acid sequence of ORF440 with that of NrtA from Synechococcus PCC 7942 as optimted by the FASTA program (Pearson and
Lipman, 1988). Vertical lines indicate aligned and identical amino acid residues.
Colons indicate conserved amino acid residue replacements. 36
ORF440 exhibited better than 28% sequence identity to NrtA over a span of 440 arnino acids.
Sequence data obtained from pBC5O for the region downstream of ORF146
(Figure 2) was initially analyzed using the BLAST program as described previously.
Approximately 1200 bp downstream of ORFI 46 was sequenced and initial sequence analysis did not reveal any regions with sequence similarity to sequences in the international databases. Visual inspection of the region downstream of ORF146 identified two potential hair-pin structures that may serve as translation terminators
(Figure 3, lower case). The first potential hair-pin has a neck of 6 bp and a loop of 8 bases. The second has a neck of 8 bp and a loop of 10 bases.
Insertional Inactivation of ORF146 and ORF440
To investigate the role of ORFI 46 in Synechococcus PCC 7942, an insertionally inactivated mutant was generated. The plasmid construct that was used to transform
Synechococcus PCC 7942 and the strategy for its construction is shown in Figure 8.
The Xbal-HinuiIl region of pBH22, containing ORF146, was subctoned into pUC19 to serve as the basis for the transformation construct. An a- cassette conferring resistance to spectinomycin was excised from pHP45n (nSpcr, Prentki and Krisch,
1984) and ligated into the Bgnl site of ORF146. The resulting plasmid, pXH::ORFI 46, was used to transform Synechococcus PCC 7942 cells. Transformants were obtained after 5 days of growth at 5% CO,, after which they were replated to fresh media. A total Figure 8. Cloning strategy for the construction of a ORFI 46 insertional inactivation construct, pXH::ORF146, for transformation and targeted mutagenesis of endogenous
ORF146 in Synechococcus PCC 7942. The Synechococcus transformation vector was constructed by subcloning a region containing ORF146, and then disrupting ORFI 46 with the 0-Spec cassette of pHP450. Shaded arrows indicate the position of open reading frarnes (ORF) and the direction of transcription of each ORF. The ampicillin resistance gene is also indicated (Ampr), as are relevant restriction enzyme sites. 1 \ Hindlll Bglll 1. Subclone Xbal-Hindll fragment in to pUC19
Xbal
2. lsolate Spcr cassette by digestion of pHP45- Omega with Barn HI and ligate tYI into Bgl II site of pXH. Barn HI and Bgl II have compatable cohesive ends that religate and form a Bst YI restriction site.
3. Select spectinornycin and ampicillin resistant transformants for plasmid isolation, and restriction analysis. Plasmid pXH::ORF146 cannot replicate in Synechococcus PCC 7942. BstYl Figure 9. Cloning strategy for the development of an ORF440 insertional inactivation construct, pP12::ORF440, for transformation and targeted mutagenesis of ORF440 in
Synechococcus PCC 7942. The Synechococcus transformation vector was constructed by subcloning a region containing ORF440, and then disrupting ORF440 with the n-
Kanr cassette of pBSLI28. Shaded arrows indicate the position of open reading frames
(ORF) and the direction of transcription of each ORE The ampicillin resistance gene is also indicated (Ampr), as are relevant restriction enzyme sites. Subclone Pstl fragment ORF440 into pUC19.
Pstl /'
\ Pst1 / Pstl 2. Isolate, and ligate an' /cassette from pBSL128 / into Clal site of pP12. 3. Select kanamycin-ampicillin resistant transforrnants for plasrnid isolation and restriction analysis. Plasmid pP12::ORF440 cannot replicate in Synechococcus PCC 7942. UGP of three transfers to fresh media were made to ensure that the intended lesion had completely segregated. Mutants were also plated to ampicillin to verify that the spectinomycin resistant mutants that were obtained resulted from double homologous recombination. Spectinomycin resistant cells only arise from recombination since the transformation vector is unable to use the replication, transcription and translation machinery of Synechococcus PCC 7942 to propagate itself. Cells able to grow on ampicillin were the result of a single recombination event that integrated the transformation vector into the genome, which maintains the wild type genomic copy of
ORFI 46. Only those ce1ls that were am picillin sensitive and spectinomycin resistant were selected for further analysis.
To determine the genotype of the mutants, Southern analysis of genomic DNA was performed (Figure 10, lanes 1 and 2). In wild type cells, genomic DNA digested with Clal and hybridized with an EcoRI-Bglll fragment of ORF146 yielded a hybridization signal corresponding to a size of approximately 5.0 kb (Figure 10, lane
2). In the ::ORFI46 mutant, designated CSI, a Clal hybridization signal of approximately 7.0 kb was obtained using the same probe (Figure 10, lane 1). The -2.0 kb increase in the size of the hybridizing DNA corresponds well with the size of the cassette, wbich is -2.0 kb. It is also apparent that only a single band exists in the lane containing DNA from CS1 , indicating that the wild type copy of ORF146 was eliminated and that the intended lesion resulted from a double recombination event.
To determine if ORF440,ORF263, and ORF289 play a role in cyanate transport and/or metabolism a mutant was generated. This mutant would also indicate if the Figure 10. Southern hybridization analysis of Synechococcus PCC 7942 CS1 and CA1 strains. Genomic DNA from CS1 and wild type cells of Synechococcus PCC 7942 was isolated, restriction enzyme digested with Clal (lane 1 and 2, respectively), electrophoresced on 0.8% agarose gels for 2 hours at 60 V, and Southern blotted.
Southern hybridization was performed at 65°C with the 187 bp EcoRI-Bglll region of
ORFI 46 (obtained from pBH22) serving as the probe. Southern hybridization analysis of CA1 and wild type cells (lanes 3 and 4, respectively) was performed essentially as described above, however genomic DNA was digested with Psd and the 687 bp Psfl-
Clal region of pP12 served as the probe. genes upstream of ORFI 46 form a functional unit. ie. an operon, along with ORFI 46.
If they are. then disrupting the upstream gene (ORF440) would result in disruption of the other genes, including ORFI 46. Disruption of ORF440 would preclude expression of downstream genes because of the design of the antibiotic resistance cassette used.
The cassette is designed such that the antibiotic resistance gene is flanked by transcription-translation stop sequences which prematurely terminate expression of sequences upstream and downstream of the cassette. A targeted mutant of this gene cluster was generated using the kanamycin resistance cassette @-Kanr) of plasmid pBSL128 (Alexeyev et al., 1995). The scheme used to generate the ORF440 insertionally inactivated mutant is outlined in Figure 9. The Psfi region of pE56, which included most of ORF440 (1 197 bp), was excised and ligated into pUC19 to create plasmid pP12. Plasmid pP12::ORF440 was constructed by ligating the n-Kanr into the
Clal site of pP12. The resulting plasmid was used to transform cells of Synechococcus
PCC 7942 as described previously. Kanamycin resistant transformants were obtained after growth for 8 days at 5% CO, and subsequently streaked out to fresh BG-11 plates supplemented with kanamycin a total of three times to ensure complete segregation of the intended Iesion.
The genotype of the ::ORF440 mutant, designated CA1, was determined using
Southern hybridization analysis (Figure 10, lanes 3 and 4). Genomic DNA from wild type Synechococcus PCC 7942 cells digested with Psn (Figure 10, lane 4) and probed with the 686 bp Psfl-Clal region of pP12, results in a hybridization signal of approximately 1200 bp. The same probe used on genomic DNA from CA1 results in 67 one strong hybridization signal at approximately 2700 bp. Both values are in agreement with what was expected for a double recombination mutant of ORF440, considering that the antibiotic resistance cassette used introduced a Pstl site into the region of interest, creating a Pstl restriction fragment of approximately 2700 bp in CA1.
Furthermore, the absence of a 1200 bp band in lane 2 (Figure 10) indicated that the wild type copy of ORF440 had been eliminated, and that the intended lesion had fully segregated.
Cyanate Dependent O, Evolution and Fluorescence Quenching in WT and CS1
Cells of Synechococcus PCC 7942
The O,-electrode provides a convenient means to measure photosynthesis by monitoring O, evolution from cells as a function of time. In the context of cyanate- dependent 0,evolution, it provides a rapid method for determining if the CS1 mutant is unable to metabolize cyanate to CO,. If the mutant is unable to generate CO, from cyanate, cyanate should not be able to support photosynthetic 0,evolution. At the same tirne it should be unable to form a C, pool, which would be reflected in an inability to quench chlorophyll a fluorescence. Prior to determining the effect of cyanate on photosynthesis and fluorescence, it was required to determine how the CS1 mutant performed photosynthetically when presented with Ci. Standing culture cells of wild type Synechococcus PCC 7942 and CS1 were given 25 mM NaCl and allowed to reach the CO2compensation point before photosynthesis was initiated by the addition of 20 68 pM Ci (Figure 1 IB). A maximum photosynthetic rate of 60 pmol O, mg'' Ch1 h-' was supported by the wild type and 67.11 pmol O, mg-' Chl h-' for CSI. While measurements of O2evolution were being made so were measurements of chlorophyll
-a fluorescence quenching. Both wild type and CS1 cells demonstrated Ci-dependent fluorescence quenching (Figure 11A) indicative of the formation of an intracellular pool of Ci. Wild type cells were able to quench fluorescence to 35.7% of Fv*, while CS1 cells quenched 35.6% of Fv*. These results indicated that the mutant cells were as photosynthetically competent as wild type cells, and supported rates of Ci-dependent photosynthetic 0, evolution, and chlorophyll a fluorescence quenching that were similar to wild type cells.
Cyanate-dependent O, evolution and chlorophyll a fluorescence quenching was examined in wild type and CS1 cells following the addition of 1 mM KOCN. Prior to the addition of KOCN, cells were given 25 mM NaCl and allowed to photosynthesize until net O, evolution ceased. The addition of 1 mM KOCN to wild type cells resulted in quenching of chlorophyll g fluorescence (Figure 11 C) and 0,evolution (Figure 11 D).
The addition of KOCN resulted in chlorophyll a fluorescence quenching to a maximum of 42.9% of F,*. Cyanate (1 mM) supported a rate of photosynthetic O, evolution of
70.1 mol O, mg-' Ch1 kt'. The addition of 1 mM KOCN to CS1 cells resulted in a minor transient in fluorescence quenching which returned to Fm*within 45 seconds. This may be due to the small amount of HCO; (< 5 PM in 1 rnM KOCN) that is present in the
KOCN as a contaminant. Subsequently, there was no further fluorescence quenching until20 pM Ci was added to the CS1 cells. The addition of Ci resulted in quenching of Figure 11. Fluorescence quenching and photosynthetic oxygen evolution in wild type
(WT, - ) and CS1 insertional inactivants (CS1 , ------) of Synechococcus PCC 7942.
Ci-dependent fluorescence quenching (A) and photosynthetic O, evolution (B) were initiated by the addition of 20 FM KHCO, to cells after they had reached the CO, compensation point. Cyanate-dependent fluorescence quenching (C) and photosynthetic 4 evolution (D) were initiated by the addition of 1 mM KOCN to cells that had reached the CO, compensation point. Cells were incubated at 30°C in a thermostatted Clarke-type electrode. Actinic light was provided at 100 pmol (photons) mQ s-' (PPFD). fluorescence to a maximum of 38.5% of Fv*. The addition of KOCN to CS1 cells also did not result in O, evolution. There was no increase in O, evolution at any time during the time course of KOCN addition. The addition of 20 pM Ci did result in a net production of O,, indicating that the cell suspension was capable of photosynthesis in spite of the presence of cyanate. These results indicate that CS1 cells are unable to support cyanate-dependent photosynthetic O, evolution or cyanate-dependent chlorophyll a fluorescence quenching, whereas wild type cells are capable of both.
Mass Spectrometric Analysis of Synechococcus PCC 7942 WT and CS1
Mass spectrometry has been used in the past to demonstrate the ability of cyanobacterial cells to deplete the extracellular medium of CO,. In the current work, mass spectrometry was used to demonstrate the presence or absence of cyanate- dependent CO, efflux from Synechococcus PCC 7942. The addition of 1 mM KOCN to high-CO, grown wild type cells (Figure 12A) resulted in a net efflux of CO, as indicated by an increase in the m/z= 44 signal from the mass spectrometer. Cyanate addition resulted in an immediate efflux of CO, to the extracellular buffer, increasing the extracellular CO, concentration to greater than 1 PM. CO, efflux continued for the entire time course, however the rate of CO, efflux decreased with time. After 10 minutes the extracellular CO, concentration reached approximately 3 PM.The addition of 1 mM KOCN to high-CO, grown CS1 cells did not result in an efflux of CO,. For the entire time course, there was no net efflux of CO, from the CS1 cells. The same set of IL
Figure 12. Mass spectrornetric rneasurement of CO2 efflux by high-CO, grown (A) and standing culture cells (B) of wild type (WT) and CA1 cells of Synechococcus PCC
7942. CO, efflux was measured using a magnetic sector mass spectrometer with a membrane inlet system that is attached to a 30°C thermostatted chamber. Cells were allowed to deplete the medium of Ci prior to the addition of 2 mM KOCN in the light
(supplied at 210 pmol (photons) mQs-' (PPFD)). Time, min
Time, min 74
experiments were performed on SCC of wild type and CS1 cells (Figure 128). A net
efflux of CO, was detected from wild type cells upon the addition of 1 mM KOCN,
however, the amount of CO2 detected was less than that detected from high-CO,
grown cells (approximately 50% less). An initial increase in CO, efflux was followed by
a gradua1 decease in the rate of efflux, which eventually levelled of at approximately
1.25 FM. The addition of 1 mM KOCN to CS1 cells did not result in an efflux of CO, at
any point over the entire time course. These results indicate that the addition of KOCN to wild type cells of Synechococcus PCC 7942 results in CO2efflux to the extracellular
buffer. This was not apparent in CS1 cells. Similar results were obtained for SCC,
however net efflux of CO, from wild type cells was less.
Cyanate-Dependent O, Evolution and Fluorescence Quenching in WT and CA1
Cells of Synechococcus PCC 7942
5'ynechococcus PCC 7942 wild type and CA1 cells were assayed for cyanate-
dependent O, evolution and fluorescence quenching using the same methodology
described previously. Standing culture cells of both wild type and CA1 Synechococcus
PCC 7942 were initially assayed for Ci-dependent O, evolution and fluorescence
quenching to ensure that they were both photosynthetically competent. Wild type cells,
in the presence of 25 mM NaCI, were allowed to deplete the buffer of contaminating Ci
until net 0, evolution ceased, at which time 20 MMCi was added (Figure 138). Addition of Ci resulted in net O, evolution, reaching a maximum rate of 50.0 pmol 4 mg'' Chl Figure 13. Fluorescence quenching and photosynthetic oxygen evolution in wild type
(WT, - ) and the CA1 insertional inactivant (CAI, ------) of Synechococcus PCC
7942.Ci-dependent fluorescence quenching (A) and photosynthetic O2 evolution (B) were initiated by the addition of 20 pM KHCO, to cells after they had reached the CO, compensation point. Cyanate-dependent fluorescence quenching (C) and photosynthetic 0,evolution (D) were initiated by the addition of 1 mM KOCN to cells that had reached the CO2 compensation point. Cells were incubated at 30°C in a thermostatted Clarke-type electrode. Actinic light was provided at 100 pmo l (photons) m" s-' (PPFD). h". The addition of 20 pM Ci to CA1 cells also resulted in net O, evolution, which attained a maximum rate of 61.8 pmol O, mg-' Chi h". Simultaneous measurements of chlorophyll a fluorescence quenching were made for each cell type (Figure 1 3A), wh ile
O, measurernents were being made. The addition of 20 pM Ci to wild type cells resulted in a rapid quenching of fluorescence, reaching a maximum of 52.0% of F;. Similarly, the addition of 20 pM Ci to CA1 cells resulted in a rapid quenching of fluorescence, reaching a maximum of 40.4% of F/.
The addition of 1 mM KOCN to wild type cells resulted in a rapid production of
O, (Figure 13D). The maximum rate of O, evolution supported by 1 mM KOCN was
54.8 pmol0, mg-' Chl h". Further addition of 20 PM Ci to wild type cells did not result in any significant increase in O, evolution. The addition of 1 mM KOCN to wild type cells also resulted in rapid quenching of fluorescence (Figure 13C). Fluorescence was quenched to a maximum of 65.8% of Fv* in wild type cells. Fluorescence was quenched an additional 5.8% (total of 71.6%) when 20 pM Ci was added to wild type cells. The addition of 1 mM KOCN to CA1 cells did not result in any significant increase in 0,evolution. The lack of 4 production was evident for the entire time course. 0, production was observed when CA1 cells were presented with 20 pm Ci. The addition of 1 mM KOCN to CA1 cells resulted in a slight increase in fluorescence quenching, reaching a maximum of 9.4% of Fv*. The addition of 20 pM Ci resulted in a rapid quenching of fluorescence, to a maximum of 45.5% of Fv*. Taken together, these results indicate that CA1 cells are unable to support cyanate-dependent photosynthetic
O, evolution or cyanate-dependent chlorophyll a fluorescence quenching, whereas wild type cells are capable of both.
Cyanate Transport in WT, CS1, and CA1 Cells of Synechococcus PCC 7942
Cyanate transport was assayed using the silicone fluid filtering centrifugation technique, with K014CNserving as the substrate. Experiments were conducted in a
Clarke-type electrode so that photosynthetic O, evolution could be monitored as the
substrate was being added. Wild type cells of Synechococcus PCC 7942 at the CO,
compensation point were administered KOl4CNto a final concentration of 2 mM. Upon
addition of the substrate, there was a rapid increase in O, evolution (data not shown).
Samples taken 5 minutes after K014CN addition accumulated 14Cipools of 27.1 mM
(SD= + 9.1, n= 8). Addition of K014CNto CS1 cells at the CO, compensation point did
not result in significant O, evolution (data not shown), as in Figure 11D. Samples taken
at 5 minutes accumulated 014CN' pools of approximately 4.1 mM (SD= 2.9, n= 8).
The CA1 mutant accumulated a 014CN- pool of 3.1 mM (SD= I1.22, n= 6) after 5
minutes of incubation with the substrate. Addition of K014CNto CA1 cells at the CO,
compensation point did not result in significant O, evolution (data not shown). Wild type
cells incubated in the dark accumulated a 14Cipool of -0.1 mM (SD= I 0.6, n= 7) after
being incubated with the substrate for 5 minutes. - - Northern Analysis of RNA Transcript Size From Wi, CS1 and CA1 Cells of
Synechococcus PCC 7942
Northern hybridization analysis of total RNA from SCC of Synechococcus PCC
7942 was used to determine the size of ORF146 and ORF440 transcripts. The entire coding sequence of ORF146 (Figure 2) served as the probe for examining its transcription. A strong hybridization signal was obtained and estimated to be approximately 0.5 kb in size (Figure 14 WT, lane 2). A weaker hybridization signal was also evident, corresponding to a size of approximately 3.9 kb. Between these two bands there was evidence of diffuse smearing. Total RNA obtained from SCC of CS1 mutants and probed with the ORF146 coding-sequence did not produce significant hybridization signais (Figure 14 CSI, lane 2). The same probe used on total RNA from
CA1 cells produced a band of medium intensity, corresponding to a size of approximately 0.5 kb (Figure 14 CA1, lane 2). There was no evidence of a band at 3.9 kb. The ORF440 coding-sequence probe produced a smeared hybridization signal when used on total RNA from wild type cells (Figure 14 WT, lane 1). The smear extends from approx. 1.4 kb to 3.9 kb. Within the smear, a band of approximately 2.0 kb was evident. The CS1 mutant gave similar results, however, the hybridization signal was not as intense and the smear did not extend as far up the lane as it did for wild type cells (Figure 14 CSI, lane 1). CA1 cells exhibit a marked absence of hybridization to the ORF440 coding sequence probe. Only very faint smearing was evident, in the
range of 1.6 to 3.4 kb (Figure 14 CA1, lane 2). Figure 14. Northern hybridization analysis of Synechococcus PCC 7942 wild type
(WT), CS1 and CA1 standing cells culture. Total RNA was isolated and treated as described earlier. Approximately 20 pg of RNA was electrophoresced under denaturing conditions in 1.2% agarose gels for 2 hours at 60 V, and blotted. Hybridizations were carried out at 60°C as were post hybridization washes. RNA contained in Lane 1 from
WT, CSI, and CA1 was probed with the 1197 bp Psil region of cynA (A). RNA contained in Lane 2 of WT, CSI, and CA1 was probed with the 465 bp EcoRI-Bglll region of cynS (S).
DISCUSSION
The work of Miller and Espie (1994) provided evidence for cyanase-like activity in the cyanobacterium Synechococcus UTEX 625. This study provides molecular and physiological evidence for the presence of a cynS-homolog in the cyanobacterium
Synechococcus PCC 7942. A 4.5 kb genomic region was isolated, sequenced and characterized. Four putative ORF's were identified based on their sequence similarity to existing genes. Three of the ORF's exhibit significant sequence identity to proteins that are members of ABC transport systems. The fourth gene (ORFI 46) was identified as an E. colCcynS hornolog, also designated cynS. To characterize cynS in
Synechococcus PCC 7942, a mutant strain was generated by means of insertional inactivation. Characterization of this strain provided evidence for the participation of cynS in cyanate metabolism. The close spatial arrangement of the genes identified in this study indicated that they may form an operon. To examine this possibility a mutant strain was generated in which the first gene (ORF440) was insertionally inactivated.
Physiological and molecular characterization of this strain provided evidence supporting the idea that these genes exist in an operon.
cynS is Responsible for Cyanate Metabolism in Synechococcus PCC 7942
The cynS gene of Synechococcus PCC 7942 was initially identified based on its sequence identity to cynS from E. coli. Insertional inactivation of cynS abolished ua
cyanate metabolism, indicating that the ORF identified was responsible for cyanate
metabolism in this cyanobacterium. Cyanate metabolism in Synechococcus PCC 7942
caused chlorophyll fluorescence quenching and photosynthetic O, evolution
indicating that the products of its breakdown are able to be used to generate a Ci pool
which can in turn support photosynthesis. Cyanate metabolism also resulted in an
efflux of CO, out of the pool, as detected by mass spectrometry. Miller and Espie
(1994) determined that the products of cyanate metabolism in Synechococcus UTEX
625 are CO, and NH,, similar to what was observed for cyanate breakdown in E. coli
(Johnson and Anderson, 1987). Given the fact that Synechococcus UTEX 625 and
Synechococcus PCC 7942 are closely related (Golden et al., 1989), it seems
reasonable to assume that cyanate is metabolized via a similar reaction in
Synechococcus PCC 7924, producing CO, and NH,.
The CS1 mutant of Synechococcus PCC 7942 possesses an insertionally
inactivated cynS gene. This strain exhibited a marked absence of cyanate metabolism.
It was unable to support cyanate-dependent chlorophyll a fluorescence quenching,
photosynthetic O, evolution or CO, efflux. It may be argued that these processes are
not operating in CS1 because of cyanate poisoning. However, before and after cyanate
addition, CS1 cells were capable of supporting Ci-dependent chlorophyll 2 fluorescence quenching and photosynthetic 0, evolution. This indicates that their
ability to generate a Ci pool, and support photosynthesis are unaffected by the
presence of cyanate or by the lesion at cynS, at least over the short term. The addition
of Ci after cyanate did not result in O, evolution greater than that expected from Ci w7 available for fixation, indicating that the absence of cyanate metabolism was not a consequence of a HCO; deficiency.
In E. col( cyanase is CO-expressedwith carbonic anhydrase (CA). Considering that the cyanase-mediated decomposition of cyanate is a HC0; dependent reaction, it seems reasonable for CA to exist in an operon with cyanase in E. coli. In the absence of CA, E. coli cells deplete what HCO,' is present without eliminating cyanate completely. In the absence of HCO;, cyanate cannot be broken down and is toxic to the cell (Kozliak et al., 1995; Guilloton et al., 1992). So how does Synechococcus PCC
7942 prevent depletion of HCO, when metabolizing cyanate? It is a well established fact that Synechococcus PCC 7942 uses its CCM to generate a large intracellular Ci pool. HCOj is a major constituent of the Ci pool and therefore, it would appear that the role of CA in E. coliis played by the CCM and the Cipool in Synechococcus PCC 7942.
In fact, CA in cyanobacteria converts HCOi to CO2 for use in photosynthesis. Cyanate metabolism produces CO2which can either equilibrate with HCO,' within the cytosol or leak out of the cell. CO, which is released from the cell can be re-transported by the
CO, transport system and in turn re-establish the Ci pool. In these terms it doesn't seem that there will be a lack of HCOifor cyanate metabolism in Synechococcus PCC
7942.
The results obtained from CS1 provide evidence for the involvement of cynS in cyanate metabolism in Synechococcus PCC 7942. It may be argued that cyanate metabolism in CS1 was not observed because the substrate for metabolism was not available. Perhaps disruption of cynS affects cyanate uptake from the extracellular 85 environment. The question of substrate availability was addressed using the silicone fluid filtering centrifugation technique. A measured 014CN' pool of approximately 4.1 mM (I 2.9, n= 8) was determined to be available after 5 minutes of incubation with
K014CN. The pool exists solely as 0î4CN' since contaminating I4Ci would be photosynthetically fixed into acid-stable products by the time the first samples were taken, since CS1 cells are fully capable of photosynthesis although they are deficient in cyanate metabolism. The absence of photosynthetic 0, evolution in the presence of a 014CN' pool indicates that cyanate metabolism was impaired in CS1 cells, further implicating cynS as the gene responsible for cyanate metabolism in Synechococcus
PCC 7942.
cynS is Expressed as Part of an Operon
The four ORF's which have been identified in this study are closely spaced with respect to each other. Although this observation in itself may not provide convincing evidence for the organization of these ORF's in an operon, the fact that the three genes upstream of cynS share identity with genes that are rnembers of ABC transport operons lends support to this idea. To investigate this possibility, an insertionaliy inactivated mutant strain (CAI) was generated that was disrupted at ORF440 (cynA). The premise for this strategy was that disruption of the upstream gene of an operon with a gene cassette that prematurely terminates transcription will essentially elim inate expression of the targeted gene and those which are downstream of it that are also members of the operon. If cynS was not a member of an operon, then cyanate metabolism should
not be affected. If cynS was a member of the operon, then its expression would be
affected and cyanate would not be metabolized.
The CA1 mutant was unable to support cyanate-dependent chlorophyll a fluorescence quenching and photosynthetic 0,evolution. These cells were capable of
supporting Ci-dependent fluorescence quenching and photosynthetic 0, evolution,
prior to, and after cyanate addition, indicating that the observations made were not a
result of cyanate poisoning. Upon the addition of Ci, no more O, was evolved than that
expected from Ci fixation, indicating that HCOi def iciency was not a contributing factor towards a lack of cyanate metabolism. The ORF's identified upstream of cynS share
sequence identity with members of ABC transport systems, suggesting that they may
in fact be responsible for cyanate transport (considering their linkage to cynS). If
cyanate was the substrate for transport, then disrupting transport would affect the
availability of cyanate for metabolism Le. none would occur. Measurements using the
silicone fluid centrifugation technique indicated that a OI4CN-pool of 3.05 mM (SD=
1.22, n= 6) existed after 5 minutes, and that during this time no significant O, evolution occurred. The absence of cyanate metabolism in the presence of substrate indicates that the lesion at cynA affected cynS expression, providing evidence that cynA, B, D, and S are linked in an operon.
To further investigate the possibility of cynS belonging to an operon that
includes cynA (and presumably cynB and cynD), Northern analysis was performed to determine the size of the cynS transcript. A cynS specific probe identified two transcripts from wild type SCC of Synechococcus PCC 7942 that differ significantly with
respect to their size and abundance. The more abundant smaller transcript was crudely
estimated to be approximately 0.5 kb. This value is what rnight be expected if cynS was transcribed on its own. The larger, less abundant, transcript was estimated to be
approximately 3.9 kb. This is what might be expected if al1 four genes are transcribed
as an operon. The identification of the larger transcript by a cynS-specific probe
provides evidence that cynS was expressed as part of a larger transcript, and that it
may be part of an operon. However, the greater abundance of the smaller sized cynS transcript suggests that cynS may also be transcribed independently of the other genes. The absence of both transcripts from CS1 cells indicates that the observed
hybridization signals are likely not the result non-specific hybridization.
A cynA specific probe identified a range of transcripts, likely indicating instability of the primary transcript. This does not appear to be unusual as similar observations
have been made for the nitrate assimilation operons of Anabeana PCC 7120 (Frias et
al., 1997) and Synechococcus PCC 7942 (Luque et al., 1994). However, of note is the fact that the largest transcript identified by the cynA probe was approximately the same size as the large transcript identified by the cynsprobe. The size similarity of these two transcripts, although not definitive, provides evidence for the existence of cynA and
cynS (and presumably cynB and cynD) in an operon.
Although it appears that cynS is part of an operon, Northern analysis of the CA1 mutant with the cynS probe identified a low abundance of cynS transcript. The absence of larger transcripts, similar to what was observed for the wild type, indicates that the - -
intended lesion effectively disrupted expression of cynA and the genes downstream of
it. However, the presence of a low abundance of what seems to be cynS transcript
indicates that cynS may still be expressed in CA1 cells. The obvious absence of
cyanate metabolism in CA1 contradicts this finding, and suggests that perhaps the CA1
cynS transcript was unable to produce a functional CynS. The absence of cyanate
metabolism because of a lack of HCOB has already been considered and found to not
be a likeiy reason. Furtherrnore, the absence of metabolism was not a consequence
of substrate unavailability since a 014CN-pool of 3 mM was measured in CA1. Taken together, these results indicate that cynS is expressed as part of an operon.
Cyanate Uptake in Synechococcus PCC 7942
Physiologica.I characterization of CA1 indicated an absence of cyanate
metabolism. Transcript analysis of the wild type and mutant strains indicated that the
genes identified in this study behave as an operon. Considering the identity of the deduced amino acid sequences of cynA, cynB, and cynD to proteins that are members
of ABC transporters, it seems reasonable to suggest that these genes code for proteins that actively transport a particular substrate. Given their linkage to cynS, the most
obvious substrate for transport would be cyanate. Although cyanate may be an obvious choice, it is likely not the natural substrate for transport. Cyanate is generally not
present in the environment but may arise from the spontaneous decomposition of urea
in aqueous solutions ( Dirnhuber and Schutz, 1948; Kemp and Kohnstam, 1956; Marier and Rose, 1964; Hagel et a/., 1971). Cyanate may also be produced intracellularl y f rom
the spontaneous decomposition of carbamyl phosphate (Allan and Jones, 1964). If the
genes identified in this study are responsible for cyanate transport, disruption of their
expression should eliminate cyanate uptake. The accumulation of a 3 mM 0g4CN'pool
in the CA1 mutant indicated that cyanate was still being taken up by these cells.
However, before refuting the involvement of these genes in cyanate transport, cyanate
uptake by other means should also be considered.
The question of cyanate transport has not been addressed in the literature, even for E. col. The cynT gene of E. coli was initially identified as a cyanate permease
(Sung and Fuchs, 1989), but later determined to be a carbonic anhydrase (Guilloton
et al., 1992). Before considering other modes of uptake, the facilitated diffusion of
cyanate will be considered first. Although the OCN- ion can itself not cross the plasma
membrane, cyanic acid (HOCN) can diffuse across the membrane. With respect to the
cyanate transport assays, the addition of 2 mM KOCN (which dissociates to K+ and
OCN') to pH 8.0 medium results in an extracellular [HOCN] of 100 nM (pK, HOCN= 3.7
(Lister, 1955)). Assuming that HOCN rapidly equilibrates across the plasma mernbrane,
the maximum intracellular [HOCN] will also be 100 nM. Cells in the light have an
intracellular pH of approximately 7.8 (Coleman and Colman, 1981; Ritchie, 1991 ),
resulting in an intracellular [OCN'] of 1.26 mM. Thus, about one-half of the observed
pool could be accounted for by facilitated diffusion.
Andher possible route for cyanate uptake is via the CO, transport system.
Cyanate (O=C=N] is a structural analog of CO, (O=C=O)that has been shown to inhibit CO, transport (Miller et a/., 1991). Initial work by Espie and Tong (unpublished)
has gone a long way in elucidating possible means of cyanate uptake in
Synechococcus UTEX 625. Several lines of evidence exist for the involvernent of the
Ci transport system in cyanate uptake. The silicone fluid filtering centrifugation technique was used to determine that ethoxyzolamide (EZ) reduces 0t4CN-uptake by
64% in Synechococcus UTEX 625 (Espie and Tong, unpublished). EZ is a known
inhibitor of active CO, transport and Na+-independent HCO,' transport (Price and
Badger, 1989; Tyrrell et aL, 1996). Reduction of total 0t4CN'uptake by EZ indicates that a proportion of gross O'CN' uptake occurs via the Ci transport system. The fact that 014CN' uptake was not cornpletely inhibited indicates that some other means of
uptake also exists. If 014CN' uptake was solely dependent on the Ci transport systern, then no uptake would be expected above the level supported by diffusion. However if
014CN' uptake was independent of the Ci transport system, essentially no reduction in
uptake, as compared to the minus- EZ control, would be expected.
Competition experiments also provided evidence for the involvernent of the Ci transport system in cyanate transport. The use of high concentrations of '*Ci caused
an initial 50% reduction in 014CN' uptake (Espie and Tong, unpublished). In addition,
12Ci caused a 64% decrease in the size of the rneasured 14C pool. This pool was
presumed to be primarily 014CN' since Ci transport was saturated by 12Ci . The
possible retransport of 014CN--derived l4CO2was reduced by dilution with 12Ci (Espie
and Tong, unpublished). The absence of complete inhibition of 014CN-uptake and I4C
pool formation by excess 12Ciindicated that 014CN- uptake can occur by some other means, but that Ci transport does contribute to 014CN-uptake. Again, if cyanate uptake
was solely dependent on Ci transport, no uptake or pool formation would be expected.
Furthermore, that uptake and pool formation occur in the presence of excess Ci
indicates that if an alternate route for cyanate uptake does exist, it is not inhibited or very poorly inhibited by Ci. If it was inhibited, then no 014CN- uptake would be expected.
To determine which component of the Ci transport system was contributing to cyanate transport, the affect of increasing concentrations of cyanate on Ci transport was examined (Espie and Tong, unpublished). Concentrations of cyanate, up to 1.5
mM, had no significant affect on Na'independent or Na'dependent HCOj transport.
At a concentration of 1.25 mM, cyanate reduced CO, transport by 50 to 60% in both
SCC and high-Ci grown cells of Synechococcus UTEX 625. The fact that increasing
concentrations of cyanate progressively reduced CO, transport indicates that cyanate
interacts with the transport mechanism in some way and in doing so, inhibits CO, transport. These results suggest that cyanate can compete with CO, for transport via the CO, transport system, which in turn accounts for inhibition of 014CN-uptake by EZ.
The fact that the two modes of HCO,' transport are not affected by cyanate, indicates that they may not be involved in cyanate uptake, and that the CO, transport component
of the Ci transport system plays a more significant role in cyanate uptake.
The work of Espie and Tong (unpublished) provides good evidence for the
involvement of the CO, transport system in cyanate uptake. Their work also provides
evidence supporting cyanate uptake by some other means. Miller and Espie (1994) YZ determined that cyanate metabolism did not occur in the dark, suggesting an energy requirement. Cyanate uptake via the CO, transporter would require light since this system is light-dependent (Miller et al, 1988). In this study wild type cells exhibited essentially no cyanate-dependent 0, evolution in the dark, nor was there any significant 014CN' uptake in the dark. The former was consistent with results obtained by Miller and Espie (1994) for Synechococcus UTEX 625, and the latter consistent with results obtained by Espie and Tong (unpublished), who obsewed essentially no uptake of 0I4CN*in the dark, or in the presence of DCMU. The reduction in 014CN-uptake was essentially the same in either case. Collectively these results indicate that cyanate uptake in Synechococcus PCC 7942 and UTEX 625 has some requirement for energy.
The decrease in uptake resulting from DCMU provides more convincing evidence for an energy dependence since a decrease in intracellular 014CN' in the dark could result from acidification of the cytosol. In the dark, the intracellular pH decreases to about 7.2
(Coleman and Colman, 1981; Ritchie, 1991). At an extracellular pH of 8.0, 100 nM of
H014CNwill exist upon the addition of 2 mM K014CN. Upon equilibration across the periplasmic membrane, 316 pM 014CN' will be present in the cytosol. Thus light stimulation of cyanate uptake may be due to alkalinization of the cytosol rather than light activated transport. Nevertheless, DCMU inhibition of cyanate uptake suggests the involvernent of energy. If the energy requirement was restricted to cyanate uptake via the CO, transport system, then levels of 014CN' uptake would be expected to be similar to those obtained using EZ. That they are not indicates that the other mode of uptake also requires energy. The identification of genes coding for an ABC transporter, that are also linked to cyanase, indicates that they rnay serve as the alternate route for cyanate uptake.
Although elimination of the putative substrate recognition gene (CA1 strain) results in pool formation, these cells likely took up cyanate by some other means, i.e. facilitated diffusion andor the CO, transporter. The CS1 mutant seemed to be a good candidate for assessing the role of the other genes in cyanate transport since it would eliminate cyanate metabolism and allow for the determination of the 014CN' pool. Determining the 014CN- pool in wild type cells is complicated by 014CN' metabolism. Cyanate metabolism yields l4CO2,which can either be fixed, form a 14Ci pool or be expelled.
14C02 that is expelled can be retransported and participate in pool formation.
Whatever the case may bel an accurate measurement of the 0I4CN' pool cannot be made since in wild type cells the measured acid-labile pool reflects 14Ci and 014CN'.
Transcript analysis of the CS1 mutant indicated a slightly reduced level of the cynABDS transcript relative to the wild type. The significance of this interaction, or even if one exists, remains to be determined. A cynS mutant that does not affect expression of the upstream genes would be desired for a more accurate assessrnent of cyanate uptake in the absence of cyanate metabolism.
Physiological Significance of Cyanate Lyase in Synechococcus PCC 7942
The physiological significance of cyanase in E. coli seems to be that of cyanate detoxification, which in turn prevents inhibition of growth (Anderson et al., IWO). The inducible nature of this system in E. coli indicates that the cells are not normally confronted with enough cyanate in the environment or intracellularly to warrant a system that is constitutively expressed at a high level. In Synechococcus PCC 7942
(and UTEX 625) cynS is expressed constitutively under photoautotrophic growth conditions, indicating that these cells are likely exposed to cyanate more so than E. coli. Therefor, they require a system that is capable of rapidly degrading cyanate before it reaches levels that are toxic. It seems unlikely that Synechococcus PCC 7942 encounters significant levels of cyanate in the environment, therefore it seems probable that cyanate must arise from an endogenous source. Expression of cynS in
Synechococcus PCC 7942 without cyanate being present in the growth media supports this possibility. The most obvious endogenous source of cyanate is carbamyl phosphate.
Carbamyl phosphate is a precursor for arginine and pyrimidine nucleotide biosynthesis (Pierard et a/., 1965). Carbamyl phosphate (CP) biosynthesis is mediated by the enzyme carbamyl phosphate synthetase (CPSase) (Mergeay et a/., 1974). This enzyme fixes CO, into glutamine resulting in the formation of glutamate and CP
(Tabita, 1987). The enzyme ornithine carbamyl transferase (OCTase) uses CP to carbamylate ornithine, in turn yielding citrulline, which is eventually used to synthesize arginine. This pathway represents an alternate route for CO, assimilation in cyanobacteria (Tabita, 1987). Although in cyanobacteria the major CO, fixation pathway is the Calvin's cycle, this alternate pathway has been proposed as a means of providing arginine for the production of cyanophycin in cyanobacteria that make use of this nitrogen storage compound (Tabita, 1987). Synechococcus species do not
produce cyanophycin (Newman et al., 1987), however, growth in the presence of high
levels of nitrogen may result in an accumulation of amino acids including arginine
(Miller and Espie, 1994). Accumulation of arginine would likely result in an
accumulation of CP which is known to spontaneously decompose to cyanate (Allens
and Jones, 1964). Such a situation may require the operation of a constitutively
expressed cyanase which can rapidly detoxify cyanate (Miller and Espie, 1994).
Although the toxic affects of cyanate on cyanobacteria remain to be elucidated, cyanate
is a known inhibitor of many enzymes including CPSase and CA. For this reason it
seems reasonable that cynS is expressed and cyanate removed.
In spite of the known toxic effects of cyanate the CS1 strain grows well and at
a rate near that of wild type cells. Therefore, it would seem that the intracellular level
of cyanate does not, in fact, rise to a level that is inhibitory to growth, and that
detoxification of cyanate may not be the sole function of the cynS gene product.
Recent work by Suzuki et al. (1996) indicates that cyanate rnay have wide
spread effects on metabolism in cyanobacteria by regulating genes that participate in
carbon and nitrogen assimilation. Cyanate (and CP) represses expression of the nitrate
assimilation operon (nirA operon) and strongly activates expression of Rubisco.
Furthermore, cyanate was shown to have a negative effect on the expression of
glutamine synthetase (glnA), which is responsible for the synthesis of glutamine, which
in turn is a precursor for the production of CP. Suzuki et al. (1996) propose that
cyanate derived from the spontaneous decomposition of CP may serve as a metabolic -- signal for the ammonium-promoted regulation of genes involved in nitrate and CO,
assimilation in Synechococcus PCC 7942. A similar situation of by-product inhibition exists in E. coli where cyanate derived from CP negatively regulates CPSase expression (Guilloton and Karst, 1987). That cyanate-mediated regulation of the nirA operon exists seems reasonable in light of the identification of cynS in Synechococcus
PCC 7942. cynS metabolizes cyanate and in doing so provides the cell with NH,.
Repressing expression of the nirA operon by cyanate reduces what might be conceived as the unnecessary production of NH,, by means of NO; uptake and its eventual
reduction to arnmonia. If cyanate can provide NH, then there is no reason for the nirA operon to be expressed at the expense of metabolic energy. However, given the tremendous capacity of Synechococcus PCC 7942 to decompose cyanate, the
likelihood that cyanate would act as a persistant intracellular signal can easily be
questioned. Obviously there is a need to measure the in vivo levels of cyanate during
metabolic shifts and at steady state.
Future Research Objectives
Transcript Analysis
This study identif ied cynS in the cyanobacterium Synechococcus PCC 7942.
Initial work identified two cynS transcripts, a large one indicative of an operon, and a
small one indicating independent transcription of cynS. Whether or not these are - - distinct transcripts remains to be determined. That the CA1 mutant appears to produce a cynS transcript while being unable to metabolize cyanate is problematic. Primer extension analysis of cynS, using primers specific for the cynS coding region and that of the potential operon transcription start site, can be used to determine the actual transcription start site. If needed, SI nuclease protection assays can also be employed to map the transcriptional start sitek of cynS.
Cyanate Transport
Determining how cyanate is taken up is a question that rernains not completely answered. Although it seems unlikely that the ABC transporter identified in the current study is used to transport cyanate, some evidence for its involvement does exist. The
0I4CN-pool declined from 4 mM in the CS1 strain to 3 mM in the CA1 strain indicating that the lesion ai cynA effected 014CN' uptake. However, it remains unclear what other modes of transport exist for cyanate uptake. The CO, transport system has been implicated along with facilitated diffusion, however cyanate uptake seems to also occur by sorne other rneans. One problem that confounds estimating cyanate uptake and pool formation is cyanate metabolism. Although the CS1 mutant is deficient in metabolism, the intended lesion may also affect expression of the upstream genes. If this is the case, then the CS1 mutant does not allow for an accurate assessment of cyanate uptake in the absence of metabolism because it may preclude uptake (in any) via the
ABC transporter identified in this study. A cynS mutant is required where expression of the upstream genes is not affected.
Using an insertional inactivation scheme that utilizes a different antibiotic resistance cassette may facilitate generation of a mutant strain that is exclusively disrupted at cynS. The cassette used in this study bears expression termination sequences flanking the antibiotic resistance gene. Premature transcription termination of the operon rnay cause instability in the remainder of the transcript and in doing so affects expression of the upstream genes. A cassette without such sequences will allow for transcription through the entire region without premature termination. Such a cassette will result in a full length transcript, from which normal translation can occur.
The position of the cassette will prevent translation of cynS but not that of itself or that of the upstrearn genes.
Once such a cynS mutant is generated, a more accurate determination of cyanate uptake and pool formation can be made. lnhibitors of CO, transport can be used to determine to what extent other modes of transport contribute to cyanate uptake. lnhibitors of other transport systems, such as those of NO; and Soi, can also be used to determine if these systems contribute to cyanate uptake. To determine if the
ABC transporter identified contributes to cyanate transport an cynAB mutant would be required. In such a mutant the components that would contribute to uptake, namely the substrate recognition domain and integral membrane domain, would be eliminated, essentially negating cyanate uptake via this pathway. However such a mutant would also require disruption of cynS so that only cyanate uptake and pool formation are measured. YY
Physiological Significance of Cyanate Lyase in Synechococcus PCC 7942
Under the growth conditions used in this study, the mutant strains CS1 and CA1
did not exhibit any obvious signs of growth inhibition indicating that the genes targeted
may code for proteins that are not essential. Cyanate has been implicated as a global
regulator of carbon and nitrogen assimilation (Suzuki et aL, 1996) and the apparent
constitutive expression of cynS indicates that cyanate levels are present at high
enough concentrations to warrant continuous cyanase activity. Long term experiments
need to be performed to determine if higher concentrations of cyanate are inhibitory to
mutant cell types. This study determined that after short exposures to cyanate (1 0
minutes) there was no apparent decrease in the ability of cells to photosynthesize, or
transport and form a Ci pool. Long term exposure to cyanate may have effects on these
processes. To determine if such affects exist, mutant cells in the presence of cyanate
will be measured with respect to their ability to photosynthesize and transport Ci. A
progressive reduction in photosynthetic output andlor Ci transport compared to wild type would indicate that cyanate affects these processes after a period of longer
exposure. Furthermore, long term growth experiments need to be performed to
determine if exogenously supplied cyanate effects the growth of the CS1 strain.
Concluding Remarks
The present study has identified four putative genes that participate in cyanate metabolism. An insertional inactivant of cynS was unable to metabolize cyanate, but did not exhibit any obvious signs of inhibition of growth, indicating that cynS is responsible for cyanate metabolism in Synechococcus PCC 7942. This study provides molecular and physiological evidence for the cyanate lyase-mediated decomposition of cyanate in the cyanobacterium Synechococcus PCC 7942. Alexeyev MF, Shokolenko INyCroughan TP (1995) lmproved antibiotic-resistance gene cassettes and omega elements for Escherichia coli vector construction and in vitro delet ionlinsertion mutagenesis. Gene 160: 63-67.
Allen CM, Jones ME (1964) Decomposition of carbamylphosphate in aqueous solutions. Biochem 3: 1238-1247.
Altschul SF, Gish W, Miller Wy Myers EW, Lipman DJ (1990) Basic local alignment search tool. J Mol Biol 215: 403-41 0.
Anderson PM (1 980) Purification and properties of the inducible enzyme cyanase.
Biochem 19: 2882-2888.
Anderson PM, Carlson JD (1975) Reversible reaction of cyanate with a sulfhydryl group at the glutamine binding site of carbamyl phosphate synthetase. Biochem 14:
3688-3694.
Anderson PM, Little RM (1986) Kinetic properties of cyanase. Biochem 25: 1621-
1626. Anderson PM, Sung Y-C, Fuchs JA (1990) The cyanase operon and cyanate
metabolism. FEMS Microbiol Rev 87: 247-252.
Anderson PM, Johnson WV, Endrizzi JA, Little RM, Korte JJ (1987) Interaction of
mono- and dianions with cyanase: evidence for apparent half-site binding. Biochem 26:
3938-3943.
Anderson PM, Johnson WV, Korte JJ, Xiong X, Sung Y-C,Fuchs JA (1988)
Reversible dissociation of active octamer cyanase to inactive dimer promoted by
alteration of the sulfhydryl group. J Biol Chem 263: 5674-5680.
Anderson PM, Korte JJ, Holcomb TA, Cho Y, Son C, Sung Y-C (1994) Formation
of intersubunit disulfide bonds and properties of the single histidine and cysteine
residues in each subunit relative to the decameric structure of cyanase. J Biol Chern
269: 15036- 1 5045.
Andrews TJ, Lorimer GH (1987) Rubisco: structure, mechanisms, and prospects for
improvement. ln: Hatch MD, Boardman NR (eds). The Biochemistry of Plants. A
comprehensive treatise. Academic Press, San Diego. Vol 10, pp 131-21 8. Ausubel FM, Brent R, Kingston RE, Moore DD, Seidman JG, Smith JA, Struhl K
(1993) Current Protocols in Molecular Biology, John Wiley & Sons, New York.
Badger MR (1987) The CO2 concentrating mechanism in aquatic phototrophs. In:
Hatch MD, Boardman NR (eds). The Biochemistry of Plants. A cornprehensive treatise.
Academic Press, San Diego. Vol 10, pp 21 9-274.
Badger MR, Andrews TJ (1982) Photosynthesis and inorganic carbon usage by the marine cyanobacterium Synechococcus sp.. Plant Physiol70: 517-523.
Badger MR, Price GD (1989) Carbonic anhydrase activity associated with the cyanobacterium Synechococcus PCC7942. Plant Physiol89: 51-60.
Badger MR, Price GD (1990) Carbon oxysulfide is an inhibitor of both CO, and HCO; uptake in the cyanobacteriurn Synechococcus PCC 7942. Plant Physiol 94: 35-39.
Bairoch A (1992) PROSITE: A dictionary of sites and patterns in proteins. Nucleic
Acids Res 20: 201 3-2018.
Bertini 1, Borghi E, Luchinat C, Scouafava A (1981) Binding sites of anions in superoxide dismutase. J Am Chern Soc 103: 7779-7783. Buch K (1960) Dissoziation der kohlensaure, gleichgenichte und puffersysteme. In:
Ruhland W (ed). Handbuch der fflanzenphysiologie. Springer-Verlag, Berlin. Vol 1 pp.
1-11.
Chin CCQ, Anderson PM, Wold F (1983) The amino acid sequence of Escherichia coli cyanase. J Biol Chern 258: 276-282.
Chollet R, Anderson PM (1978) Cyanate modification of lysyl residues in the catalytic subunits of tobacco ribulose diphosphate carboxylase. Biochrn Biophys Acta 525: 455-
467.
Coleman JR, Colman B (1981) lnorganic carbon accumulation and photosynthesis in a blue-green algae as a function of external pH. Plant Physiol67: 917-921.
Coleman JR, Seemann JR, Berry JA (1982) RuBP carboxylase in carboxysomes of blue-green algae. Carnegie lnst Wash Year Book 81: 83-87.
Codd GA, Marsden WJN (1984) The carboxysome (polyhedral bodies) of autotrophic prokaryotes. Bi01 Rev 59: 389-422.
Codd GA (1988) Carboxysomes and ribulose bisphosphate carboxylaseloxygenase. - -- In: Rose AH, Tempest DW (eds). Advances in Mirobial physiology. Academic Press,
San Diego. Vol 29, 115-1 64.
Crotty CM, Tyrrell PN, Espie GS (1994) Quenching of chlorophyll g fluorescence in response to Na+-dependentHCO; transport-mediated accumilation of inorganic carbon in the cyanobacterium Synechococcus UTEX 625. Plant Physiol104: 785-791.
Dirnhuber P, Schutz F (1948) The isomeric transformation of urea into ammonium cyanate in aqueous solutions. Biochem J 42: 628-632.
Engberg P, Lindskog S (1984) Effect of pH and inhibitors on cobalt II carbonic anhydrase from bovine skeletal muscle. FEBS Letters 170: 326-336.
Espie GS, Canvin DT (1987) Evidence for Na+-independent HCO, uptake by the cyanobacterium Synechococcus leopoliensis. Plant Physiol 84: 125-130.
Espie GS, Kandasamy RA (1 992) Na+-independentHCO, transport and accumulation in the cyanobacterium Synechococcus UTEX 625. Plant Physiol98: 560-568.
Espie GS, Kandasamy RA (1994) Monesin inhibition of Na'-dependent HCOj transport distinguishes it from Na+-independent HCOj transport and provides evidence for Na+/HCOj symport in the cyanobacterium Synechococcus UTEX 625. Plant Physiol
104: 1419-1428.
Espie CS, Miller AG, Canvin DT (1988) Characterization of the Na+-requirementin cyanobacterial photosynthesis. Plant Physiol 88: 757-763.
Espie OS, Miller AG, Canvin DT (1989) Selective and reversible inhibition of active
CO2transport by hydrogen sulfide in a cyanobacterium. Plant Physiol91: 387-394
Espie GS, Miller AG, Canvin DT (1991) High affinity transport of CO, in the cyanobacterium Synechococcus UTEX 625. Plant Physiol97: 943-953.
Espie GS, Miller AG, Kandasamy RA, Canvin DT (1991) Active HCO, transport in cyanobacteria. Can J Bot 69: 936-944.
Frias JE, Flores E, Herrero A (1997) Nitrate assimilation gene cluster from the heterocyst-forming cyanobacterium Anabeana sp. Strain PCC 71 20. J Bacteriol 179:
477-486.
Forster D, Goodgame DML (1965) Vibrational spectra of pseudo-halide complexes
II. Planar and octahedral isocyanate complexes. J Chem Soc 1286-1289. Golden SS, Nalty MS, Cho DC (1989) Genetic relationship of two highly studied
Synechococcus strains designated Anacystis nidulans. J Bacteriol 171:24-29.
Guilloton M, Hargreaves AB (1972) Hydrolyse du cyanate par une bactérie du genre
Flavobacterium. C R Acad Sc Paris 275: 1827-1 830.
Guilloton M, Karst F (1987a) Isolation and characterization of Escherischia coli
mutants lacking inducible cyanase. J Gen Microbiol 133: 645-653.
Guilloton M, Karst F (1987b) Cyanate specifically inhibits arginine biosynthesis in
Esherischia cok a case of by-product inhibition. J Gen Microbiol 133: 655-665.
Guilloton MB, Korte JJ, Lamblin AF, Fuchs JA, Anderson PM (1992) Carbonic
anhydrase in Escherichia cok a product of the cyn operon. J Biol Chem 267: 3731-
3734.
Hagel P, Gerding JJT, Fieggen W, Bloemendal H (1971) Cyanate formation in
solutions of urea. Biochm Biophys Acta 243: 366-373.
Hofmann K, Stoffel W (1993) TMbase - A database of membrane spanning proteins segments. Biol. Chem. Hoppe-Seyler 347: 166 Johnson and Anderson (1987) Bicarbonate is a recycling substrate for cyanase. J
Biol Chem 262: 9021-9025.
Kaneko T, Tanaka A, Sato S, Kotani Hy Sazuka T, Miyajima N, Sugiuri M, Tabata
S (1995) Sequence analysis of the genome of the unicellular cyanobacterium
Synechocystis sp. strain PCC6803. 1. Sequence features in the 1 Mb region from map positions 64% to 92% of the genome. DNA Res 2: 153-166.
Kaplan A¶ Badger MR, Berry JA (1 980) Photosynthesis and the intracellular inorganic carbon pool in the blue-green alga Anabaena variabilis: response to external CO, concentration. Planta 149: 2 19-226.
Kemp IA, Kohnstam G (1956) The decomposition of inorganic cyanates in water. J
Chem Soc 191: 900-91 1.
Kuhlemeier CJ, van Arkel GA (1987) Host-vector systems for gene cloning in cyanobacteria. Methods Enzymol l53:199-215.
Kim KH, Honzatko RB, Little RB, Anderson PM (1987) Preliminary x-ray crystallographic study of cyanase from Escherichia coli. J Mol Biol 198: 137- 138. 1 UY
Kotani H, Kaneko A, Matsubayashi T, Sato S, Sugiura My Tabata S (1994) A physical rnap of the genome of a unicellular cyanobacterium Synechocysfis sp. strain
PCC6803. DNA Res 1 : 303-307.
Kotani H, Tanaka A, Kaneko TySato S, Sugiura M, Tabata S (1 995) Assignment of
82 known genes and gene clusters on the genome of the unicellular cyanobacterium
Synechocystis sp. strain PCC 6803. DNA Res 2: 133-142.
Kozliak El, Fuchs JA, Guilloton MB, Anderson PM (1995) Role of bicarbonate/C02 in the inhibition of Esherishia coli growth by cyanate. J Bacteriol 177: 3213-321 9.
Kunz DA, Nagappan O (1989) Cyanase-mediated utilization of cyanate in
Pseudomonas fluorescens NClB 11764. Appl Environ Microbiol55: 256-258.
Kyte J, Doolittle RF (1982) A simple rnethod for displaying the hydrophobic character of a protein. J Mol Biol 157: 210-213.
Laudenbach DE, Grossman AR (1 991 ) Characterization and mutagenesis of sulfur- regulated genes in a cyanobacteriurn: Evidence for function in sulfate transport. J
Bacteriol 173: 2739-2750. I IV
Lister MW (1955) Some observations on cyanic acid and cyanates. Can J Biochem 33:
426-440.
Little RM, Anderson PM (1987) Structural properties of cyanase: denaturation, renaturation, and role of sulfhydryls and oligomeric structure in catalytic activity. J Bi01
Chem 262: 10120-101 26.
Luque 1, Flores E, Herrero A (1994) Molecular mechanism for the operation of nitrogen control in cyanobacteria. EMBO J 13: 2862-2869.
Mackinney TA (1941) Absorption of light by chlorophyll solutions. J Biol Chem 140:
315-322.
Marier JR, Rose D (1964) Determination of cyanate, and a study of its accumilation in aqueous solutions of urea. Anal Biochem 7: 304-314.
McKay RML, Gibbs SP, Espie OS (1993) Effect of dissolved inorganic carbon on the expression of carboxysomes, localization of Rubisco and the mode of inorganic carbon transport in cells of the cyanobacterium Synechococcus UTEX 625. Arch Microbiol
159: 2 1-29. ... Mergeay M, Gigot D, Beckman J, Glansdorff N, Piérard A (1974) Physiology and genetics of carbamylphosphate synthesis in Escherichia coli KI2. Mol Gen Genet 133:
299-316.
Miller AG (1990) lnorganic carbon transport and accumulation in cyanobacteria. In:
Codd G A (ed). Autotrophic microbiology and one-carbon metabolism. Kluwer
Academic Publishers, Dordrecht. pp 25-53.
Miller AG, Canvin DT (1987a) The quenching of chlorophyll a fluorescence as a consequence of the transport of inorganic carbon by the cyanobacteriurn
Synechococcus UTEX 625. Biochm Biophys Acta 894: 407-413.
Miller AG, Canvin DT (1987b) Na+-stimulationof photosynthesis in the cyanobacterium
Synechococcus UTEX 625 grown on high levels of inorganic carbon. Plant Physiol84:
118-124.
Miller AG, Canvin DT (1989) Glycoaldehyde inhibits CO, fixation in the cyanobacterium Synechococcus UTEX 625 without inhibiting the accumilation of inorganic carbon or the associated quenching of chlorophyll a fluroescence. Plant
Physiol 91 : 1044- 1049.
Miller AG, Colman B (1980a) Evidence for HCO; transport by the blue-green alga (cyanobacterium) Coccochloris peniocystis. Plant Physiol65: 397-402.
Miller AG, Colman B (1980b) Active transport and accumulation of bicarbonate by a unicellular cyanobacterium. J Bacteriol 143: 1253-1259.
Miller AG, Esple GS (1994) Photosynthetic metabolism of cyanate by the cyanobacterium Synechococcus UTEX 625. Arch Microbiol (1994) 162: 151 -157.
Miller AG, Espie OS, Canvin DT (1988) Active transport of CO2by the cyanobacteriurn
Synechococcus UTEX 625. Plant Physiol86: 677-683.
Miller AG, Espie GS, Canvin DT (1989) Use of carbon oxysulfide, a structural analog of CO2,to study active transport of CO2 in the cyanobacterium Synechococcus UTEX Newman J, Wyman MyCarr NG (1987) Absence of the nitrogen reserve polymer cyanophycin from the marine Synechococcus species. FEMS Lett 44: 221-224.
Ogawa T, Togasaki RK (1988) Carbonyl sulfide: an inhibitor of inorganic carbon transport in cyanobacteria. Plant Physiol88: 800-804.
Otwinowski Z, Anderson PM, Fuchs JA, Sigler PB, Joachimiak A (1991)
Proceedings of the IV International Meeting on Crystallization of Biological
Macromolecules. p 54 August 18-24 Freiburg, Germany.
Pearson WR, Lipman DJ (1988) lmproved tools for biological sequence comparison.
Proc Natl Acad Sci USA 85: 2444-2448.
Piérard A, Glansdorff N, Mergeay M, Wiame JM (1965) Control of the biosynthesis of carbamyl phosphate in Escherichia coli. J Mol Biot 14: 23-36.
Prentki P, Krisch HM (1984) In vitro insertional mutagenesis with a selectable DNA fragment. Gene 29: 303-313.
Price GD, Badger MR (1989) Ethoxyzolarnide inhibition of CO, uptake in the cyanobacteriurn Synechococcus PCC7942 without apparent inhibition of interna1 carbonic an hydrase activity. Plant Physiol 89: 37-43.
Price GD, Badger MR (1991) Evidence for the role of carboxysomes in the cyanobacterial CO,-concentrat hg mechanism. Can J Bot 69: 963-973.
Reinhold L, Menekhern Z, Kaplan A (1989) A quantitative model for inorganic carbon fluxes and photosynthesis in cyanobacteria. Plant Physiol Biochem 27: 945-954.
Reinhold L, Kosloff R, Kaplan A (1991) A model for inorganic carbon fluxes and photosynthesis in cyanobacterial carboxysornes. Can J Bot 69: 984-988.
Ritchie RJ (1991) Membrane potential and pH control in the cyanobacterium
Synechococcus R-2 (Anacystis nidulans) PCC 7942. Plant Physiol137: 409-418.
Saiki RK (1990) Amplification of genomic DNA. In: lnnis MA, Gelfand DH, Sninsky,
White TJ (eds). PCR Protocols: A Guide to Methods and Applications. Academic Press,
Inc., San Deigo. pp. 13-20.
Sambrook J, Fritsch EF, Maniatis T (1989) Molecular Cloning: A Laboratory Manual.
Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 1 la
Shelp BJ, Canvin DT (1984) Evidence for bicarbonate accumulation by Anacystis
nidulans. Can J Bot 62: 1398-1403.
Schreiber U, Schliwa U, Bilger W (1986) Continuous recording of photochernical and
non-photochernical chlorophyll fluoprescence quenching with a new type of modulation f luororneter. Photosynth Res IO: 51-62.
Stark GR (1972) Modification of proteins with cyanate. Methods Enzymol 258: 579-
584.
Strommer J, Gregerson R, Vayda M (1993) Isolation and characterization of plant
mRNA. In: Glick BR, Thompson GE (eds). Methods in Plant Molecular Biology and
Biotechnology. CRC Press, Boca Raton. pp. 49-65.
Sung Y-C, Fuchs JA (1988) Characterization of the cyn operon in Escherichia coli
KI2. J Bacteriol263: 14769-14775.
Sung Y-C, Fuchs JA (1989) Identification and characterization of a cyanate permease
in Escherichia coli K- 12. J Bacteriol 171 : 4674-4678.
Sung Y-C, Fuchs JA (1992) The Escherichia col/ K-12 cyn operon is positively regulated by a member of the IysRfamily. J Bacteriol 174: 3645-3650.
Sung Y-C, Parsell D, Anderson PM, Fuchs JA (1987a) Identification, mapping, and cloning of the gene encoding cyanase in Escherichia coli K-12. J Bacteriol 169: 2639-
2642.
Sung Y-C, Anderson PM, Fuchs JA (1987b) Characterization of high-level expression and sequencing of the Escherichia coli K- 1 2 cynS gene encoding cyanase. J Bacteriol
169: 5224-5230.
Suzuki 1, Sugiyama T, Omata T (1996) Regulation by cyanate of the genes involved in carbon and nitrogen assimilation in the cyanobacteriurn Synechococcus sp. strain
PCC 7942. J Bacteriol 178: 2688-2694.
Tabita FR (1 987) Carbon dioxide fixation and its regulation in cyanobacteria. In: Fay
P, Van Baalen C (eds). The Cyanobacteria. Elsevier Biomedical Press, Amsterdam. pp.
96- 11 7.
Taussig A (1 960) Effect of carbamylphosphate and cyanate on Escherischia coli. Can
J Microbiol 6: 619-629. 11/
Tyrrell PNy Kandasamy RA, Crotty CM, Espie GS (1996) Ethoxyzolamide differentially inhibits CO, uptake and Na+-independent and Na+-dependent HCOj uptake in the cyanobacterium Synechococcus sp. UTEX 625. Plant Physiol 112: 79-88.
Volokita M, Zenvirth D, Kaplan A, Reinhold L (1984) Nature of the inorganic carbon species actively taken up by the cyanobacterium Anabaena variabilis. Plant Physiol 76:
599-602.
Yu J-W, Price GD, Song L, Badger MR (1992) Isolation of a putative carboxysomal carbonic anhydrase gene from the cyanobacterium Synechocystis PCC7942. Plant
Physiol 100: 794-800. APPLIED 2 IMAGE. lnc 1653 East Main Street ,-.- Rochester, NY 14609 USA ------Phone: 71 61482-0300 ------FM: 7161288-5969
Q 1993, Applied Image, Inc.. All Rights Resewed