Biochemical and physiological characterisation of Deg/HtrA proteases in Synechocystis sp. PCC 6803
Dissertation
zur Erlangung des akademischen Grades des Doktors der Naturwissenschaften (Dr. rer. nat.)
an der
Universität Konstanz
Mathematisch-Naturwissenschaftliche Sektion Fachbereich Biologie
vorgelegt von Manuela Perthold
Tag der mündlichen Prüfung: 25. April 2014 1. Referent: Dr. Dietmar Funck
2. Referent: apl. Prof. Dr. Hendrik Küpper
Abstract 3
Zusammenfassung 4
I. Introduction 6
1. Cyanobacteria 6
2. Proteases 16
3. Deg/HtrA proteases 18
4. Methods to study protein-protein interactions 30
5. Aims of this work 33
II. Results and Discussion 34
1. Synechocystis sp. PCC 6803 Deg/HtrAΔPDZ mutants were unable to form higher-order oligomers 34
2. Characterisation of the E. colidegP knockout strain JW0157 41
3. Methods for substrate identification were optimised regarding their use in cyanobacteria 46
4. The effects of combined salt stress and deletion of HhoA on photosynthesis- relevant proteins and pigments 56
III. Conclusion and Perspectives 67
IV. Materials and Methods 69
1. Cyanobacterial strains and culture conditions 69
2. DNA isolation from Synechocystis sp. PCC 6803 69
3. Construction of the Synechocystis sp. PCC 6803 ΔhhoA knockout mutant 69
4. Verification of the cyanobacterial knockout mutant ΔhhoA 70
5. Determination of cell density 70
6. Salt stress conditions 70
7. Cell densities and Chlorophyll concentration 70
8. Isolation of native protein fractions from Synechocystis sp. PCC 6803 cells 71
9. Pigment fluorescence kinetic measurements 71
10. Statistical analysis 73
11. Plasmid construction for heterologous expression of Synechocystis sp. PCC 6803 HhoA in E. coli 73
12. Plasmids for TAPtag purification 73
13. Plasmids for oligomerisation experiments 74
14. Heterologous expression of Deg/HtrA constructs in E. coli 74
15. Purification of overexpressed proteins 74
16. Size exclusion chromatography 74
17. Terminal amine isotope labelling of substrates 76
18. Pulldown assay 76
19. SDS-PAGES 76
20. Silver staining analysis 77
21. Immunoblot analysis 77
22. Isolation of genomic DNA from different E. coli strains 78
23. Verification of E. coli JW0157 degP knockout 78
24. Growth ability of an E. coli degP null strain JW0157 in comparison to WT strain BW25113 78
25. PCR primers 79
26. Vectors 82
V. Acknowledgements 83
VI. Appendix 84
1. Figures 84
2. Tables 85
3. List of Abbreviations 85
VII. Literature 87
Abstract
Proteases catalyse the hydrolysis of peptide bonds and participate in many cellular processes, such as maturation, degradation and turnover of proteins. Deg/HtrA proteases are ATP-independent serine endoproteases that can be found in almost every organism. Besides the protease domain with a catalytically active serine, they possess up to four PDZ domains mediating protein-protein interactions, substrate identification and complex formation. To this date, relatively little is known about the physiological interaction partners of the Deg/HtrA proteases. The three Deg/HtrA proteases HhoA, HtrA, and HhoB in Synechocystis sp. PCC 6803 are of special interest regarding their interaction partners, as these proteases combine features of their plant and bacterial homologues and are thought to be involved in responses to abiotic stresses and the turnover of the PSII reaction centre protein D1. This study focused on the role of the cyanobacterial Deg/HtrA proteases in the short-term response to elevated salt concentrations and particularly on changes in the levels of photosynthesis-related proteins and fluorescence kinetics. Furthermore, state-of-the-art methods for substrate identification were tested for their practicability in cyanobacteria and the biochemical characterisation of the three Synechocystis sp. PCC 6803 enzymes was extended by determining the role of the PDZ domain in the oligomerisation process.
Investigating the potential function of HhoA in the cyanobacterial salt stress response by the analysis of a ΔhhoA knockout mutant, it was found that the strain showed no impairments regarding its APC protein levels, but lower PsbO and RuBisCO levels than under control conditions. Furthermore, RuBisCO was faster restored than in WT and the growth rate of the mutant were significantly lower than in WT after 4 h salt stress. In parallel the ΔhhoA chlorophyll content per cell significantly decreased less than in the WT during the salt stress. It might be that HhoA mediates degradation or processing of one of the enzymes involved in chlorophyll degradation or synthesis and further negatively regulates pathways providing energy for cell division. In ΔhhoA, the D1 level was unaffected by salt stress, whereas in WT, D1 was completely absent after 6 h of stress. Therefore, HhoA might be involved in degradation of D1 and PsbO, at least under salt stress conditions. It was further shown by fluorescence kinetics that HhoA deletion had no impact on overall photosynthetic performance and PSII repair, even when the cells were exposed to high salt concentrations. This led to the hypothesis that HhoA is not involved in the regulation of photosynthetic processes, but in subsequent cellular salt stress adaptation mechanisms.
Several approaches to identify substrates or interaction partners of the Synechocystis sp. PCC 6803 Deg/HtrA proteases, including genetic and biochemical tools, were pursued. As part of this study, a pulldown assay was optimised with focus on a more specific binding of the bait protease HhoA to a cobalt-chelate column. Furthermore, plasmids that include a tandem affinity purification tag (TAPtag) and which were specifically designed for subsequent introduction into the Synechocystis sp. PCC 6803 genome via homologous recombination were constructed. These plasmids will be useful for co- purification of protease-interaction partner(s) complexes formed under in vivo conditions. In addition, with preparations of Synechocystis sp. PCC 6803 WT and ΔhhoA soluble proteomes, the terminal amine isotope labelling of substrates (TAILS) was tested with special interest on handling.
Binding of substrates induced the formation of large oligomers in Deg/HtrA proteases from E. coli and A.
thaliana, and the PDZ domains played a crucial role in this process. We were able to demonstrate that the single PDZ domain in the Synechocystis sp. PCC 6803 Deg/HtrA proteases facilitates trimerisation and is essential for the formation of complexes larger than trimers. Furthermore, we demonstrated that trimerisation is dependent on the pH value.
Zusammenfassung
Proteasen katalysieren die Spaltung von Peptidbindungen durch Hydrolyse und sind an vielen zellulären Prozessen beteiligt, wie zum Beispiel an Reifung, Abbau und Umsatz von Proteinen. Die Deg/HtrA- Proteasen sind ATP-unabhängige Endoproteasen, die in fast jedem Organismus vorkommen. Sie besitzen neben einer Proteasendomäne mit einem katalytisch aktiven Serin zusätzlich noch bis zu vier PDZ-Domänen, die für die Vermittlung von Protein-Protein-Interaktionen sowie Substratidentifikation und Komplexbildung zuständig sind. Relativ wenig ist bisher bekannt über potentielle Interaktionspartner der Deg/HtrA-Proteasen. In diesem Zusammenhang ist die Identifikation von Interaktionspartnern der drei Vertreter in Synechocystis sp. PCC 6803, HhoA, HtrA und HhoB, von besonderem Interesse, da diese Proteasen sowohl bakterielle als auch pflanzliche Merkmale aufweisen und möglicherweise eine Rolle in der Antwort auf abiotischen Stress spielen, sowie am Umsatz des D1- Proteins im Kern des PSII-Reaktionszentrums beteiligt sein könnten. Der Fokus dieser Arbeit lag auf der Beteiligung der cyanobakteriellen Deg/HtrA-Proteasen an der Kurzzeit-Antwort auf erhöhte Salzkonzentrationen, vor allem in Bezug auf Photosynthese-relevante Proteine und Fluoreszenzkinetik.
Weiterhin wurden neueste Techniken zur Substratidentifikation auf ihre Anwendbarkeit in Cyanobakterien getestet und die biochemische Charakterisierung der drei Synechocystis sp. PCC 6803 Deg/HtrA-Proteasen wurde mit der Bestimmung der PDZ-Domänen-Funktion bei der Oligomerisierung erweitert.
Die Untersuchung der möglichen Funktion von HhoA während der cyanobakteriellen Salzstressantwort mit Hilfe eines ΔhhoA-Mutanten zeigte keine Beeinträchtigung der Proteinexpression von Allophycocyanin in diesem Bakterienstamm.
Allerdings wurden die Konzentrationen an PsbO und RuBisCO deutlich niedriger unter Salzstress im Vergleich zur Kontrolle. Die RuBisCO-Expression erreichte im Mutanten schneller wieder die Ausgangskonzentration als im WT und nach 4 h Stressdauer war die Wachstumsrate in der Mutante deutlich niedriger als im Wildtyp, während die Chlorophyllkonzentration über die Zeit im Vergleich zum WT weniger stark reduziert wurde. Dies könnte für eine Beteiligung von HhoA an Abbau oder Reifung eines der Chlorophyll-abbauenden oder -synthetisierenden Enzyme sprechen. Weiterhin könnte HhoA als negativer Regulator in energieliefernden Prozessen für die Zellteilung beteiligt sein. Aufgrund der stark reduzierten D1-Proteinkonzentrationen im WT im Vergleich zu den nicht beeinträchtigten D1- Konzentrationen in ΔhhoA kann eine Beteiligung von HhoA am D1-Abbau unter Salzstress nicht ausgeschlossen werden.
Zusätzlich wurde über Fluoreszenzkinetik-Messungen gezeigt, dass das Fehlen von HhoA weder Auswirkungen auf die Gesamt-Photosyntheseleistung noch auf die Reparatur von PSII hat, auch wenn die Zellen hohen Salzkonzentrationen ausgesetzt sind. Stattdessen wird eine indirekte Beteiligung von HhoA an der Regulation desChlorophyll-Stoffwechsels und der Zellwandsynthese vermutet.
Verschiedene Methoden zur Identifizierung von Interaktionspartnern wurden getestet, wobei entweder genetische oder biochemische Ansätze zum Einsatz kamen. Im Rahmen dieser Arbeit wurden Plasmide gefertigt, die nach der Einbringung ins Synechocystis sp. PCC 6803-Genom über homologe Rekombination für zweistufige Protein-Affinitätsreinigungen verwendet werden können. Weiterhin wurde die einstufige Pulldown-Methode mit Fokus auf die Bindung von HhoA als Köderprotein an eine Kobaltchelat-Säule verbessert, um in Zukunft mit den Deg/HtrA-Proteasen assoziierte Proteine biochemisch untersuchen zu können. Außerdem wurde die Methode der Isotopenmarkierung von endständigen Aminen in Substraten (TAILS) im Hinblick auf ihre Anwendbarkeit in Cyanobakterien
getestet. Dafür wurden lösliche Proteine aus Synechocystis sp. PCC 6803 Wildtyp und ΔhhoA-Mutante isoliert.
Für die Substratbindung sind Protein-Protein-Wechselwirkungen notwendig, die in den gut charakterisierten Deg/HtrA-Proteasen aus E. coli und Arabidopsis hauptsächlich von PDZ-Domänen vermittelt werden. Eine weitere Aufgabe der PDZ-Domänen ist die Bildung großer Homo-Oligomere. In dieser Arbeit konnte gezeigt werden, dass die PDZ-Domänen in den Synechocystis sp. PCC 6803 Deg/HtrA-Proteasen die Trimerisierung fördern und für die Bildung von Komplexen größer als Trimere essentiell sind. Weiterhin wurde eine pH-Abhängigkeit der Trimerisierung gezeigt.
I. Introduction
1. Cyanobacteria
1.1. Cyanobacterial physiology
Cyanobacteria belong to the evolutionary most ancient organisms on earth and are thought to be more than 3.5 billion years old. They can be found in almost all habitats including hot springs, glaciers and desert areas. Cyanobacteria form a large group of gram-negative bacteria that are all capable of oxygenic photosynthesis [Kotani, H et al. 1998]. They are also considered to have contributed to the development of earth’s oxygenic atmosphere by producing molecular oxygen during photosynthesis.
According to the endosymbiotic theory, primary plastids evolved by uptake of a cyanobacterium into a heterotrophic eukaryotic host cell 1-2 billion years ago in a process called primary endosymbiosis (reviewed in [Kutschera, U et al. 2005]). Such primary plastids are present in glaucophytes, red algae, green algae and plants [Cavalier-Smith, T 2000]. Following primary endosymbiosis, most of the endosymbionts’ genes were transferred by horizontal gene transfer to the host cell nuclear genome, thereby transforming the endosymbiont into a semi-autonomous organelle with a largely reduced residual genome [Keeling, P et al. 2008]. Further features that provide evidence for the endosymbiotic events are e.g. the prokaryotic nature of the transcription and translation machinery in plastids and the double-membrane system of the chloroplast that remained of the cyanobacterial ancestor (reviewed in [Kutschera, U et al. 2005]). Thus, cyanobacteria are directly related to plastids in other photosynthetic organisms.
The most remarkable similarity between cyanobacteria and higher plant chloroplasts, further confirming the evolutionary relationship, is that the photosynthetic electron flow is basically the same [Richter, G 1998]. Both use water as electron donor for photosynthesis and are thus clearly distinguishable from purple and green photosynthetic bacteria that use e.g. sulphur as electron donor.
In cyanobacteria and plants, light is absorbed by antennae complexes and the energy is further transferred to photosystem II (PSII), where it excites chlorophyll molecules bound to the PSII core proteins D1 and D2. The excitation energy is converted into chemical energy by electron transfer to the primary acceptor phaeophytin and next to the secondary acceptor, the plastoquinone (PQ) pool. The oxidised reaction centre chlorophylls are re-reduced by electrons from the lumenal oxygen-evolving complex (OEC), where water is dissected into protons and molecular oxygen. Repeated oxidation and reduction processes of PQ create a proton gradient over the thylakoid membrane and an electron flow to the cytochrome b6/f complex (Cyt.b6/f). This complex transfers the electrons to a lumenal soluble electron carrier thereby reducing it. In plants, this lumenal carrier is the copper-containing plastocyanin.
Depending on the species and the copper availability, cyanobacteria either possess plastocyanin or cytochrome c553 (Cyt.c553; [Vermaas, W 2001]). The protons transported into the lumen by Cyt.b6/f are further used for adenosine-5’ triphosphate (ATP) synthesis, whereas the electrons are transferred to photosystem I (PSI) and are finally transferred to ferredoxin, a soluble carrier on the stromal side of the thylakoids, or, in case of cyanobacteria, on the cytoplasmic side. Ferredoxin transmits the electrons to the nicotinamide adenine dinucleotide phosphate (NADP+) reductase that produces NADPH for further use in carbon fixation or by other redox-reactive enzymes. An ATP synthase uses the energy stored in the proton gradient that is built up by both OEC and Cyt.b6/f to synthesise ATP in the stroma or cytosol of chloroplasts or cyanobacteria, respectively.
Despite the identical electron flow, the structure of the photosynthetic apparatus differs between cyanobacteria and higher plants. The photosynthetic complexes are located in the thylakoids, an endomembrane system that can be found in plastids and cyanobacteria. In chloroplasts, the thylakoids form a network of stroma lamellae and grana stacks [Gunning, B et al. 1975], whereas in cyanobacteria
and many algae, the thylakoids only form one long layer of stroma lamellae. The lamellae are filled with the thylakoid lumen, where the OEC and soluble electron carriers involved in photosynthesis can be found.
The tubular thylakoids in cyanobacteria provide much more space than grana stacks for the antenna complexes, the so-called phycobilisomes.They are membrane-associated rods of soluble phycobiliproteins that are covalently linked to a bilin chromophore via thioether bonds. Three phycobiliproteins are known in cyanobacteria, namely phycoerythrin (PE), phycocyanin (PC), and allophycocyanin (APC) and four types of bilins can be present: phycocyanobilin, phycoerythrobilin, phycourobilin, and phycoviolobilin [MacColl, R 1998]. All bilins are open-chained tetrapyrrols containing varying numbers of conjugated double bonds. The double bond system is used to absorb light energy that can be subsequently transferred from the outer phycobiliproteins, PE and PC, to the core APC and further on to chlorophyll molecules associated with PSII. As the number of conjugated double bonds decreases descending from the outside, the energy can be efficiently transferred towards the photosystems, which was called the “funnel effect” [Gantt, E 1977]. The wavelengths poorly absorbed by chlorophyll are covered by the phycobiliproteins, thus cyanobacterial phycobilisomes additionally close the “green gap” of chlorophyll.
In a structural context PE and PC are usually organised in two to six rods connected to one APC cylinder.
Each cylinder contains four trimers of the APC α/β subunits and one APC subunit contains one α- polypeptide and one β-polypeptide. In total, there are three APC cylinders with overall 12 trimers present in one phycobilisome (Figure 1).
Figure 1:Schematic structure of cyanobacterial phycobilisomes
The left picture shows the whole phycobilisome attached to the thylakoid membrane and the individual phycobiliproteins in their defined order within the antenna complex. The right picture shows a detailed composition of the phycobilisome core and rod. The core contains APC cylinders and is attached to the thylakoid membrane and thus to PSII via linker proteins. Each cylinder is composed of four APC trimers (α3β3). One trimer consists of three monomers with one α-polypeptide and one β-polypeptide, respectively. The outer proteins PC and PE are organised in up to six rods connected to one core cylinder. (pictures taken from [MacColl, R 1998]).
Another unique feature of cyanobacteria is the co-localisation of photosynthetic apparatus and respiratory chain in the thylakoid membrane. They share the PQ pool, the Cyt.b6/f complex and plastocyanin (sometimes Cyt.c553). The remaining components of the respiratory chain, namely type-1 NADPH dehydrogenase, succinate dehydrogenase, and terminal oxidase are located in the cytoplasmic membrane [Schmetterer, G 1994].
Cyanobacterial thylakoid membranes are not laterally heterogeneous, which means the photosynthetic complexes are equally distributed all over the membrane. This is in striking contrast to higher plant thylakoids, where PSII is located in the grana and PSI together with the ATP synthase is found in the stroma lamellae and the non-stacked edge regions of the grana. Another major difference between the photosynthetic apparatus of cyanobacteria and higher plants is that in cyanobacteria the stoichiometry of PSI and PSII is larger than one, whereas in higher plants the ratio is typically close to one. It is suggested that the high PSI to PSII ratio in cyanobacteria contributes to an oxidised PQ pool under high light (HL) conditions, important to reduce photodamage [Andersson, B et al. 1996].
Although cyanobacteria are classified as gram-negative bacteria, their cell wall combines features of both gram-positive and gram-negative bacteria. For example their peptidoglycan layer is thicker than that of gram-negative bacteria and contains polysaccharides more similar to those in gram-positive bacteria [Jürgens, U et al. 1986]. However, typical components of the gram-positive peptidoglycan layer, like teichoic acid, L-lysine, and diaminopimelic acid are missing in cyanobacterial cell walls [Hoiczyk, E et al. 2000]. The outer membrane contains features that are usually not present in gram-negative bacteria like carotenoids [Resch, C et al. 1983] and unusual fatty acids like β-hydroxypalmitic acid [Schrader, M et al. 1981]. The cell envelope is surrounded by 1) S-layer-forming glycoproteins, which function as protective shell and are involved in cell adhesion and recognition [Sleytr, U et al. 1996] and 2) carbohydrates that are produced to protect the membrane structure [Reeds, R et al. 1984]. Another feature that relates cyanobacteria and gram-negative bacteria is the presence of a periplasmic space, which is bordered by the cytoplasmic membrane and the outer membrane. The periplasmic space is filled with the periplasm, an aqueous compartment with jellylike texture containing enzymes, binding and transport proteins.
The most obvious distinguishing mark to eukaryotes is the absence of a nucleus in cyanobacteria. They rather contain a nucleoid composed of supercoiled deoxyribonucleic acid (DNA), which includes between 2000 and 6000 protein coding open reading frames (ORF) and is accompanied by a number of small cryptic plasmids that harbour potential protein-coding genes with yet unknown functions (http://www.ncbi.nlm.nih.gov/genome/?term=cyanobacteria). Nevertheless, the main genome codes for all proteins essential for maintenance of cellular processes. Sequence comparisons demonstrated high homology of those cyanobacterial genes to corresponding plant and green algae genes. The much simpler genome structure as well as the relationship to chloroplasts makes cyanobacteria perfect model organisms for the study of plant-like oxygenic photosynthesis and responses to abiotic stresses [Glatz, A et al. 1999; Kotani, H et al. 1998].
1.2. Synechocystis sp. PCC 6803 as model organism
Four Synechocystis culture substrains (Pasteur Culture Collection PCC, Glucose Tolerant GT, ATCC and
“Kazusa”) are used for laboratory purposes nowadays. They all derived from the fresh-water strain Berkeley 6803 [Stanier, R et al. 1971] but show different phenotypes.
The cyanobacterium most commonly used as model organism is Synechocystis sp. PCC 6803, which cannot fix nitrogen but is able to grow heterotrophically or mixotrophically when supplied with glucose
[Rippka, R et al. 1979]. Complete genome sequencing was performed in 1996 [Kaneko, T et al. 1996] and revealed 3167 ORFs covering 87% of the total genome. The first comparison of the Synechocystis sp. PCC 6803 genome to those of other photosynthetic organisms in 2001 revealed that 45% of the genes are identical or at least similar to known genes in other photosynthetic organisms, whereas 44% show no similarity to any known gene [Ikeuchi, M et al. 2001].
128 Synechocystis sp. PCC6803 genes can be assigned to functions in photosynthesis, although some plant genes like psaG (PSI subunit G) and psbQ (PSII subunit Q) are not present in Synechocystis sp. PCC 6803 (http://genome.microbedb.jp/cyanobase/Synechocystis). At least 80 genes code for two- component systems involved in signal transduction, 147 genes code for proteins required for general regulatory functions and 92 genes encode peptidases [Kaneko, T et al. 1996];
http://merops.sanger.ac.uk). This categorisation makes it relatively simple to identify at least the functional category of a newly found protein. The specific function can be investigated by using knockout mutants or loss-of-function mutants, which can be generated by integration of foreign DNA into the Synechocystis sp. PCC 6803 genome via double-homologous recombination [Grigorieva, G et al.
1982]. Heterologous expression in Escherichia coli (E. coli) and subsequent purification of proteins is another method to study protein function in vitro [Kaneko, T et al. 1997].
1.2.1 Synechocystis sp. PCC 6803 and its response to salt stress
Cyanobacteria living in habitats with changing salinity (e.g. tidal river estuaries) must be able to adapt to high salt concentrations. Although the first reactions of the organisms are responses to overcome salt stress, also fine-tuned molecular adaptation mechanisms have evolved, which are best studied in moderately halotolerant strains as Synechocystis sp. PCC 6803 and 6147 and in fresh-water strains as Synechococcus sp. PCC 7942 and 6301 [Fulda, S et al. 1999]. In Synechocystis sp. PCC 6803, which is able to grow at salinities up to 1.8 M NaCl [Joset, F et al. 1996], the salt-stress reaction and subsequent adaptation includes five major steps ([Hagemann, M 2011]; Figure 2):
Phase 1 is characterised by a sudden loss of water and soluble substances from the cyanobacterial cell by osmosis, leading to a reduction in cell size. Phase 1 continues for several ms and is followed by the 20 to 60 min-lasting phase 2. During phase 2, Na+ and Cl- ions passively enter the cell and increase the osmotic cell potential [Reeds, R et al. 1985]. When Na+ and Cl- ions reach the equilibrium with the medium, also the osmotic potential is raised above that of the medium, so that water re-enters the cell and the cell volume is restored. However, the high NaCl concentration inside the cell remains and inhibits cellular and metabolic functions, especially photosynthesis, transcription and translation processes [Marin, K et al. 2004] mainly by the toxic effect of high Na+ concentrations [Maathuis, F et al.
1999]. Therefore, in phase 3, Na+ is exchanged for K+ [Reeds, R et al. 1985] within 60 min to 2 h after salt stress implementation. K+ is thought to stabilise and define the three-dimensional structures of proteins [Maathuis, F et al. 1999]. The exchange systems include Na+/H+ antiporters as well as the monovalent cation/proton antiporter system (Mrp system) and the K+ transport system (Ktr system; [Matsuda, N et al. 2004]). The Ktr system is not dependent on ATP but on the presence of Na+ and on a high membrane potential [Matsuda, N et al. 2004]. The Mrp system is a secondary active transporter that is driven by distinct primary proton pumps [Swartz, T et al. 2005]. The exchange is followed by the synthesis and accumulation of compatible solutes. Compatible solutes are organic, low-molecular weight compounds that are highly soluble in water and do not interfere with other cellular components [Yancey, P et al.
1982]. The main compatible solute in Synechocystis sp. PCC 6803 is glucosylglycerol (GG), but also sucrose, trehalose, and glycine-betaine can be accumulated [Joset, F et al. 1996]. In summary, pre- existing processes and enzymes are activated in phases 1 to 3 to overcome immediate salt stress challenges [Kanesaki, Y et al. 2002; Marin, K et al. 2004].
The following phase 4 lasts for 3 to 8 h. It re-establishes regular gene expression and cellular function to the original level and helps the cell to acclimate to the elevated salt concentrations. In this phase, Na+ is almost completely exchanged for compatible osmolytes leading to restoration of turgor and water potential. Photosynthetic activity increases, growth and cell division are resumed. The final phase 5 is reached as early as 24 h after salt stress implementation. In this phase, the cells contain low internal ion concentrations but high levels of osmoprotectants to both protect against and adapt to the external salt concentration. Recent studies confirmed the single phases by checking the transcription state of the genes involved in the salt stress response [Marin, K et al. 2004].
Figure 2: Five-step adaptation to high external salt concentrations in Synechocystis sp. PCC 6803
The physiological changes are shown schematically in the upper part of the picture and again itemised in the lower part of the picture. The x-axis shows the time the cyanobacterium needs to fully adapt to high NaCl concentrations in the surrounding medium. The y-axis shows the levels of each adaptation process compared to the original level.
In summary, pre-existing processes and enzymes are activated in phases 1 to 3 to overcome immediate salt stress challenges. In phases 4 and 5, long-term adaptation to elevated salt concentration occurs (picture taken from [Hagemann, M 2011]). GG: Glucosylglycerol
Specific two-component systems are involved in perception of increasing salt concentrations [Shoumskaya, M et al. 2005]. Several transmembrane histidine kinases (Hik10, Hik16, Hik33, and Hik34) function as signal receptors, which sense changes in external NaCl levels. Rising ion concentrations outside the cell activate one of the Hik proteins, which in turn phosphorylates its adjacent response regulator (Rre). The Rre on the other hand enhances or lowers the expression of downstream genes.
The Hik two-component systems regulate the expression of around 70% of the stress-influenced genes in Synechocystis sp. PCC 6803. Currently, three response regulators, Rre3, Rre31 and Rre17, are known to take part in the signal transduction triggered by high salt concentrations [Shoumskaya, M et al. 2005].
Among the stress-influenced proteins, most are up-regulated at the transcriptional level, whereas only a few are down-regulated [Marin, K et al. 2004].The up-regulated genes and the corresponding proteins can be classified into four groups: salt-stress specific proteins, general stress proteins, basic carbon
metabolism proteins and unidentified proteins [Fulda, S et al. 2006; Kanesaki, Y et al. 2002]. Genes induced specifically by enhanced NaCl concentrations are for instance those coding for the Ktr system, which contributes to the Na+/K+ exchange, and the electron carrier flavodoxin (isiB, sll0248; [Fulda, S et al. 1995]). General stress proteins are e.g. GroEL1 (60 kDa chaperonin 1, slr2076), GroEL2 (60 kDa chaperonin 2, sll0416) and DnaK2 (sll0170), which are all chaperones involved in protection of proteins under stress conditions. Transcription of the GG biosynthesis pathway genes and the general σ70 stress factors is also enhanced under stress [Kanesaki, Y et al. 2002]. As carbon fixation and growth are reduced in salt-stressed cells, there must be a yet unknown reason for the up-regulation of e.g.
transketolase (tktA, sll1070) and fructose-1,6-bisphosphatase (fbpII, slr0952; [Fulda, S et al. 2006]). The largest group of proteins found to be increasingly expressed contains proteins with yet unidentified function(s).A few genes are down-regulated upon salt stress implementation, most of them coding for components of the photosynthetic apparatus and respiratory chain [Marin, K et al. 2004]. This is in line with the reduced photosynthetic activity of PSII demonstrated experimentally by [Allakhverdiev, S et al.
2002].
The OEC is also partially inhibited not because of a lower gene expression, but due to the high Na+ content in the thylakoid lumen. The Na+ ions are thought to disturb binding of the loosely attached OEC subunits to PSII and thus prevent the oxidation of water to molecular oxygen and electrons [Allakhverdiev, S et al. 2008]. A similar effect is considered to prevent binding of the soluble electron carriers plastocyanin (petE, sll0199) and Cyt.c553 (petJ, sll1796) to PSI, which decreases the rate of electrons transported to and from PSI [Allakhverdiev, S et al. 2000]. If salt stress was applied only for a short time, then both effects can be restored by water influx. Long-termed salt stress (> 5 h) leads to a dissociation of the catalytic manganese cluster of the OEC and thereby to an irreversible inhibition of photosynthesis [Allakhverdiev, S et al. 2008].
The decreased electron flow in combination with an unchanged light absorption would lead to an increase in production of reactive oxygen species (ROS), which contribute to membrane impairment, impacts on DNA and RNA as well as protein damage [Cabiscol, E et al. 2000]. On the one hand, an accumulation of ROS is minimised by the degradation of phycobilisome components under salt stress conditions [Kanesaki, Y et al. 2002]. Particularly linker proteins like apcE(slr0335) and apcF (slr1459) as well as subunits of APC (apcA, slr2067;apcB, slr1986) and PC (cpcC, sll1580; cpcB, sll1577) are down- regulated [Kanesaki, Y et al. 2002]. Simultaneously, phycobilisome degradation proteins of the nblA- group are expressed more strongly [Marin, K et al. 2004]. On the other hand, ROS production is reduced by increasing the expression of defence proteins like the water-soluble carotenoid protein (slr1963), superoxid dismutase (sodB, slr1516; [Kanesaki, Y et al. 2002] and the peroxiredoxin-like protein (sll1621;
[Fulda, S et al. 2006]).
ROS are particularly harmful to membrane lipids, as they cause lipid peroxidation of the polyunsaturated fatty acids. This modification provokes a decrease in membrane fluidity and disturbs binding of proteins to the lipid bilayer [Cabiscol, E et al. 2000]. It was found that in Synechococcus sp. PCC 6311, under salt stress conditions most membrane lipids are saturated [Huflejt, M et al. 1990], possibly to prevent unwanted peroxidation and membrane disintegration. Another mechanism of protection against ROS is to thicken the cell wall when cyanobacteria are exposed to high NaCl concentrations. This was e.g.
shown for Synechocystis sp. PCC 6803, where the expression of periplasmic proteins that are involved in cell wall synthesis and alteration is enhanced under high salt conditions [Fulda, S et al. 2000; Pandhal, J et al. 2008]. A thickened cell wall might establish a stronger diffusion barrier and thus decrease the influx of inorganic ions into the periplasm.
Especially the cyanobacterial periplasmic space is vulnerable to high salt levels, as it does not contain outer membrane transporters to actively export ions. Therefore, the periplasm directly responds to high
external salt concentrations, which can damage proteins. Periplasmic proteases to degrade salt- damaged proteins are e.g. the members of the Deg/HtrA protease family, which are also considered to act as chaperones. However, other proteases and chaperones assigned to the cytoplasm or thylakoid lumen might also be involved in defence against salt stress in the periplasm.
1.3. Methods to analyse regulation of cellular processes in Synechocystis sp. PCC 6803
1.3.1 Functional genome analysis
A powerful tool to analyse regulation of cellular processes in cyanobacteria is the genetic modification of their genome. Mutations can be introduced physically by exposure to ultraviolet (UV) light, chemically with e.g. methyl sulphonates [Herdman, M et al. 1972] or biologically by gene transfer, which can be performed by transduction, conjugation or transformation. The method of plasmid-mediated conjugation requires a plasmid that carries a mobilisation element for transfer of the desired DNA fragments, an origin of replication (ORI) and, if necessary, selectable marker regions. Subsequently, the desired fragment is transferred from the donor plasmid into the recipient cyanobacterial genome along with the ORI and the selection marker. Furthermore, replicating and non-replicating plasmid vectors (reviewed in [Thiel, T 1994]) can be used to either express an exogenous gene or transport DNA into the cyanobacterium for further use in transposon mutagenesis or homologous recombination. With transposons, the mutation is inserted randomly into one of the up to ten copies of the cyanobacterial chromosome [Herdman, M et al. 1979]. Therefore, transposon mutagenesis can be applied to initially search for mutants. Cyanobacteria, however, tend to disintegrate the transposon which is why stable integration of a particular gene is often achieved by homologous recombination [Williams, J et al. 1983].
During homologous recombination, exogenous DNA is incorporated stably into the host genome. This method is also used to test whether an effect is only caused by the introduced mutation or by spontaneous mutation elsewhere in the genome. Synechocystis sp. PCC 6803 is one of the cyanobacteria that are naturally transformable by homologous recombination [Grigorieva, G et al. 1982].
1.3.2 Analysis of photosynthetic activity by chlorophyll fluorescence measurement
In phototrophic organisms, light energy absorbed by the reaction centres is used for photosynthesis (photochemistry) to subsequently produce ATP and NADPH, which enable carbon fixation. Any excess of absorbed energy is simultaneously dissipated as heat in a process called non-photochemical quenching (NPQ) or emitted as fluorescence with a wavelength longer than the one of the initially absorbed light.
As fourth route, the absorbed energy can be transferred from one pigment molecule to the next, as it occurs e.g. in phycobilisomes of cyanobacteria. All four processes are linked in competition, which means that any change in efficiency of one parameter will influence the yield of the other three. Hence, determination of the photosynthetic capacity by measuring chlorophyll fluorescence provides information about the efficiency of heat dissipation, photochemistry and electron transport from one pigment to another. In general, fluorescence yield is highest, when the rates of NPQ and photochemistry are low.
The relationship between photosynthetic electron flow and chlorophyll fluorescence was first described by Kautsky and co-workers [Kautsky, H et al. 1960], who illuminated dark-adapted leaves and observed both an increase in chlorophyll fluorescence and changes in CO2 assimilation. This “Kautsky effect”
basically describes the transient reduction of all components of the photosynthetic electron transport chain in the initial phase after illumination, prior to their gradual re-oxidation by the onset of carbon fixation and concomitant NADP+ regeneration. The bi-phasic kinetic of the initial rise in fluorescence
intensity after light absorption by the antennae and energy transfer to the PSII reaction centre is caused by the primary quinone electron acceptor QA of PSII. QA can only accept one electron and has to pass it to the plastoquinone QB bound to PSII before receiving another electron. While QA is reduced, the photosynthetic reaction centre of PSII is closed, which leads to reduced photochemical efficiency and thus to an increased fluorescence emission. After QA has transferred its electron to QB and becomes oxidised, electron charge separation in PSII is again possible and the reaction centre is open. The chlorophyll fluorescence yield decreases correspondingly, which is termed photochemical quenching (qP; [Duysens, L et al. 1963]). Simultaneously, the NPQ rate increases until steady-state levels of qP and NPQ are reached.The real photosynthetic performance can only be determined by distinguishing between qP and NPQ.
However, eliminating NPQ contribution to fluorescence quenching is not possible. Therefore, qP is usually reduced close to zero by transiently closing all PSII reaction centres with a short, highly intensive and saturating light flash [Bradbury, M et al. 1981; Quick, W et al. 1984]. This method prevents an increase in NPQ and does not influence the long-term efficiency of photosynthesis. Thus, after switching on the measuring light, the minimal level of fluorescence (F0), in which QA is fully oxidised and all functional reaction centres are open, can be measured. The measuring light is a weak modulated measuring beam with an intensity of 0.1 µmol m-2 s-1photosynthetically active photon flux density (PPFD) that does not drive photosynthesis.
The maximum fluorescence Fm in the dark-adapted state (F0m) can be measured by applying a saturating flash of light. The following transfer form dark to light by applying actinic light (AL), which drives photosynthesis, progressively closes the PSII reaction centres and thus increases chlorophyll fluorescence. Subsequently, after a few minutes of illumination, the fluorescence level decreases by quenching. The NPQ participation to quenching is described by the parameter (F0m – F’m)/F’m. AL is interrupted by consecutive saturating light flashes that allow measuring the maximum fluorescence in the light (F’m). The steady-state fluorescence yield immediately prior to the flash (Ft) and the removal of actinic light after the flash (F’0) are further parameters that can be measured (Figure 3).
Figure 3: Overview of the different parameters measured to determine the influence of photochemical and non-photochemical quenching on chlorophyll fluorescence emission from PSII MB: Measuring beam; SP: Saturating light pulse; AL:
Actinic light; F0: Minimal fluorescence level; Fm0
: Maximum fluorescence in a dark-adapted cell; Fm’:
Maximum fluorescence in the light; Ft: Steady-state fluorescence immediately prior to the saturating light flash; F0’: Minimal fluorescence in a light- adapted cell (Picture taken from [Maxwell, K et al.
2000]).
The overall efficiency of PSII photochemistry is measured by ΦPSII, which describes the proportion of the light that is absorbed by PSII chlorophyll molecules and actually used in photochemistry [Genty, B et al.
1989]. Thus, when the amount of absorbed photons and the distribution between PSII and PSI is known, ΦPSII can be used to calculate the rate of linear electron flow and its relationship to carbon fixation, but
is strongly influenced by stress conditions due to photorespiration or the occurrence of cyclic electron transport [Fryer, M et al. 1998]. The maximum efficiency of PSII in the dark-adapted state can be characterised by Fv/Fm with Fv being calculated as Fm – F0. Thus, Fv/Fm is a measure for the quantum efficiency, if all PSII centres were open and QA maximally oxidised. In addition, Fv/Fm is used to assess damage to PSII reaction centres and in particular the occurrence of photoinhibition.
When using white light for excitation, the determination of F0 in cyanobacteria depends strongly on the cellular phycobiliprotein content and the degree of coupling to PSII reaction centres, because phycobiliproteins emit fluorescence that partially overlaps with chlorophyll fluorescence [Lee, T et al.
1994]. Because chlorophyll fluorescence is usually measured by a blue light-emitting diode (LED;
[Kolbowski, J et al. 1995]), the contribution of phycobiliproteins to F0 can be neglected and does not influence Fv in chlorophyll fluorescence measurements.
Changes in the variable PSII fluorescence result in changes of the parameter NPQ, which in plants basically reflects heat dissipation from PSII and its antennae [Adams, W et al. 1993; Horton, P et al.
1996; Krause, G et al. 1982]. In cyanobacteria on the contrary, NPQ mostly reflects changes in the distribution of the excitation energy between PSII and PSI dependent on phycobilisome state transitions [Campbell, D et al. 1996].
State transitions are the movement of antenna complexes between PSII and PSI and they constitute a way to regulate both the efficiency of electron transport between the two photosystems as well as activities of PSII and PSI in response to changing environmental conditions [Yu, L et al. 1993]. Both plants and cyanobacteria are able to change the position of their antenna complexes, which is triggered in plants by the phosphorylation state of the light-harvesting complex II (LHCII). If PSII is over-excited and the PQ pool is strongly reduced, LHCII becomes phosphorylated and migrates to PSI, whereas an over- excitation of PSI leads to LHCII dephosphorylation and subsequent movement of LHCII to PSII [Allen, J 1992; Bennet, J 1991].
In cyanobacteria, the redox state of QA regulates the position of the phycobilisomes [Mullineaux, C et al.
1990]. If PSI is overexcited, QA becomes oxidised and the phycobilisomes detach from PSI and localise to PSII (state I). Over-excitation of PSII or dark respiration [Mullineaux, C et al. 1986] closes PSII reaction centres and reduces QA. The phycobilisomes become associated with PSI (state II). PSII absorbance and fluorescence decrease in state II, whereas PSI fluorescence increases [Mullineaux, C 1992]. Since respiratory electron chain and photosynthetic electron chain use the same electron carriers, state II transition can be driven by both [Mullineaux, C et al. 1986]. State II is reflected by low variable fluorescence levels and high NPQ levels. State I transitions are characterised by higher fluorescence yields and lower NPQ levels due to partial oxidation of the electron transport chain by PSI [Campbell, D et al. 1998]. Thus, the position of the phycobilisomes also influences the redox state of the photosynthetic electron chain.
There are three proposed models for the mechanism of cyanobacterial state transition. The first model by Allen and Holmes suggests that the phycobilisomes change position, when the adjacent photosystem migrates through the membrane, taking advantage of the membrane fluidity [Allen, J et al. 1986; El Bissati, K et al. 2000]. Poly-unsaturated fatty acids promote membrane fluidity and thus photosystem/phycobilisome movement. Furthermore, over-reduction of the PQ pool in light changes the phosphorylation state of especially APC as core protein and triggers movement [Allen, J et al. 1985;
Sanders, C et al. 1986]. On the other hand, the electrostatic repulsion between the phosphorylation and the negatively charged membrane phospholipids might cause antenna movement from PSII to PSI.
Dephosphorylation thus will cause movement from PSI to PSII.
The second model [Murata, N 1969] suggests a “spill-over” of energy from PSII chlorophyll molecules to PSI chlorophyll molecules and vice versa, which are supposed to trigger phycobilisome movement [Biggins, J et al. 1989]. The authors hypothesized that the energy transfer between the photosystems is not regulated by phycobilisome movement but by changing the rate of energy transfer from PSII chlorophyll to PSI chlorophyll. The higher the PSII excitation energy amount, the more energy is transferred to PSI (state 2). In contrast, a low amount of excitation energy will decrease the transfer rate to PSI (small spill-over, state 1), because the mostly oxidised state of QA favours the use of excitation energy within PSII. The mechanism might be controlled by the distance between PSII and PSI and regulated by the relative activities of both photosystems, although this still remains to be clarified.
However, both models cannot explain the redistribution of the chlorophyll excitation energy in the absence of phycobilisome redistribution. Therefore, McConnell and co-workers proposed a combination of both models [McConnell, M et al. 2002]. The regulation of the energy transfer from the phycobilisome core to PSII is combined with regulation of the energy spill-over from the PSII core protein CP47 to PSI. The authors suggested that already small movements of PSI change the distribution of excitation energy absorbed by chlorophyll and phycobilisomes towards a more balanced state.
Nevertheless, the exact mechanism of state transitions in cyanobacteria is still not defined.
Another mechanism to regulate the energy supply in particular to PSII is either the detachment of the phycobilisomes from the thylakoid membrane or decoupling of single proteins from the phycobilisome core. Both processes decrease the energetic flow from the phycobilisomes to the PSII reaction centre [Tamary, E et al. 2012]. Subsequently, chlorophyll molecules receive fewer excitons, which reduce the risk for formation of 3chlorophyll and also ROS.
Analysis of the fluorescence emission of PSII chlorophyll, APC and PC in Synechocystis sp. PCC 6803 revealed an immediate increase in the fluorescence level after phycobilisome decoupling triggered by strong monochromatic or white light. This might arise from transient phycobilisome fluorescence quenching, which would be generated by strong light-induced pigment over-excitation, and further generation of local heat transients within the antennae complexes. These will cause modifications in the thermo-sensitive proteins that link the antennae to the photosystems, which leads to the dissociation of the phycobilisomes from the membrane. Thus, the decoupling mechanism is proposed to occur under prolonged stress conditions, when maintenance of photosynthesis is required but energy transfer is hazardous [Tamary, E et al. 2012].
Such decoupling of phycobilisome subunits was reported for cyanobacterial species like Synechocystis sp. PCC 6803 and Thermosynechococcus elongatus as well as for the unicellular red alga Porphyridium cruentum grown under HL conditions [Liu, L et al. 2009; Tamary, E et al. 2012; Yang, S et al. 2007].
Furthermore, oxidative stress induced phycobilisome decoupling in the marine filamentous cyanobacterium Trichodesmium erythraeum [Küpper, H et al. 2004]. Hence, the decoupling mechanism seems to be a general mechanism to reduce the electron flow to PSII in response to environmental stresses. Although antennae dissociation in response to high salt concentrations was not shown yet, it might also occur in order to protect PSII against ROS formation. The dissociated phycobiliproteins might be degraded, which will lead to a decrease in the overall size of the phycobilisomes leading to reduced capacity for light absorption. Thus, chlorophyll over-excitation and also photodamage are prevented.
2. Proteases
2.1 Protease functions
The removal of signal sequences, degradation of misfolded or damaged proteins and turnover of intact proteins that are not needed anymore (e.g. cyclins in the cell cycle) is catalysed by proteases. Proteases hydrolyse peptide bonds and thus irreversibly modify other proteins post-translationally. To prevent constant and unwanted degradation of proteins, protease activity has to be tightly regulated. Some enzymes are highly specific for a distinct substrate and are thus limited by its accessibility. More unspecific proteases are restricted to a specific cell compartment and/or require specific targeting of substrates. Furthermore, proteolysis can be regulated through substrate-induced conformational changes or zymogen activation. Natural inhibitors turn off protease activity reversibly or irreversibly by binding to the substrate-specificity pockets [Bode, W et al. 2000].
The main protease function is to constantly monitor the folding states of proteins and to maintain cellular homeostasis [Bukau, B et al. 2006; Wickner, S et al. 1999]. Proteases take major part in modifying receptors, transcription factors or kinases to alter their function. Additionally proteases are involved in degradation of proteins that are misfolded and/or non-functional due to e. g. gene mutations, transcriptional or translational errors [Ellis, R et al. 2006], loss of co-factors [Wickner, S et al.
1999] or stress conditions. Misfolding exposes hydrophobic patches that are normally buried within the native protein. These patches tend to bind each other, thus leading to protein aggregation and subsequent interference with normal cellular metabolism [Bukau, B et al. 2006; Ellis, R et al. 2006].
Chaperone binding to hydrophobic areas triggers protein refolding and rescue, helping proteins to reach or regain their native state and supporting complex assembly [Gottesman, S et al. 1997].Usually, chaperone binding and the attempt to refold proteins is the first step after protein misfolding occurs.
Only if refolding is not possible, the proteins are degraded by proteases. If both chaperone and protease activities fail, protein aggregates are formed that may interfere with cellular processes or deplete the cell of amino acids for translation and can cause severe illnesses like Alzheimer’s or Parkinson’s disease [Soto, C et al. 2006].
Among the most prominent proteases in eukaryotes is the ubiquitin/26S proteasome system, which is for instance involved in regulation of cell differentiation or transcription processes. The 26S proteasome is an ATP-dependent threonine protease that degrades proteins marked with polyubiquitin [Glickman, M et al. 2002]. Regulation of the proteasome system mainly occurs by the enzymatic cascade that is necessary to attach ubiquitin chains to lysine residues in target proteins. In bacteria and organelles with bacterial origin, such as chloroplasts and mitochondria, the proteasome complex is not present. Thus, other ATP-dependent enzymes like the caseinolytic protease (Clp), FtsH and Lon proteases are used for removal of damaged proteins [Adam, Z et al. 2006; Gottesman, S 2003].
2.2 Protease classification
The huge number and diversity of proteases in all living organisms has prompted several efforts to establish reasonable and practical systems to classify proteases into various subgroups. This chapter describes the classification of proteases defined at the peptidase database MEROPS (http://merops.sanger.ac.uk; [Rawlings, N et al. 2009]) and according to the International Union of Biochemistry and Molecular Biology (IUBMB; http://www.chem.qmul.ac.uk/iubmb/).
2.2.1 MEROPS classification according to the cleavage sites
All proteases cleave their substrates in a specific manner. In general, endopeptidases cleave inside the polypeptide chain and exopeptidases cut the protein at one of its termini. Exopeptidases can be further divided according to the number of amino acids they remove and on which side cleavage occurs. A special case of exopeptidases are omega-peptidases, which act on terminal residues that contain isopeptide bonds. Endopeptidases are mainly involved in limited proteolysis, i.e. removal of signal or transit peptides as well as maturation of precursor proteins but also cleave proteins into fragments that can be further processed by exopeptidases. An example of endopeptidases that create protein fragments are the Deg/HtrA proteases (see chapter 3).
2.2.2 MEROPS classification according to the chemical mechanism
The chemical mechanism defines the “catalytic type” of the enzyme, which in turn describes the chemical group mediating peptide bond hydrolysis. Normally, the amino acid that contains this group is called the catalytically active amino acid residue. So far, seven protease types are known, and six of them are further specified into serine (S), cysteine (C), aspartic (A), metallo (M), threonine (T), or glutamic (G) proteases [Hartley, B 1960]. In every case, a nucleophilic attack occurs, where a new bond is formed between the nucleophile and the substrate (electrophile) by transfer of an electron pair. The nucleophile can either be the reactive group of an amino acid side chain (S-, T-, C-types) or an activated water molecule (A-, M-type). Metallo-proteases may contain cobalt, manganese or zinc ions that coordinate and activate a water molecule as well as amino acid side chains as electrophiles. The sixth group, glutamic proteases, may have glutamate or glutamic acid as catalytically active residue. Proteases that contain more than one catalytically active amino acid are referred to as P-type proteases. An eighth group contains proteases with unknown catalytic type (U).
2.2.3 MEROPS classification according to molecular structure and homology
In the MEROPS database, proteases are grouped into families and those in turn into clans. This system is based on [Rawlings, N et al. 1993], who investigated how far the catalytic activities have diverged from one another during evolution and in contrast, how enzymes with different evolutionary background have acquired similar mechanisms. Proteases can be assigned into the same family, if the similarity of their amino acid sequences indicates a common ancestor. Therefore, especially the proteolytically active domain sequence has to be highly homologous, as this should be the most conserved part of a protease.
Each family is designated with the same letter as used for the catalytic type, since the catalytically active amino acid lies within the protease domain. The letter is followed by a unique number, so each enzyme family can clearly be identified. Within a family, individual enzymes are identified by a three-digit code that can contain both letters and numbers. The families can be further grouped into clans, if they share a common ancestor. When this cannot be deduced anymore from the primary structure, the division into clans occurs by comparison of the proteins’ three-dimensional structures. If such a structure is not available, arrangements of the catalytic domains as well as the amino acid sequences around those are looked at. A clan is specified with the letter for the catalytic type or with “P” for clans with families of several catalytic types, followed by an arbitrary letter.
2.2.4 IUBMB classification according to the chemical function
In contrast to MEROPS, IUBMB categorises enzymes primarily according to the chemical function they fulfil. That means, chymotrypsin e.g. would be a serine endoprotease in MEROPS and in IUBMB it would be hydrolase acting on peptide bonds inside a polypeptide chain with serine in the catalytic centre. The
Enzyme Commission (EC) numbering can be misleading as it groups enzymes in one class that may not be related in terms of structure or sequence. Nevertheless, there are seven EC classes described:
All enzymes oxidising their substrates are named oxidoreductases (EC class 1). Enzymes that transfer chemical groups from a donor to an acceptor are called transferases (EC class 2). Lyases (EC class 4) catalyse cleavage of molecule bonds not by hydrolysis or oxidation but by other reactions. Changes within a molecule are carried out by isomerases (EC class 5) and ligases connect two molecules (EC class 6). Class EC 3 combines enzymes that hydrolyse various bonds. Proteases belong to the subgroup EC 3.4, because they hydrolyse peptide bonds.
3. Deg/HtrA proteases 3.1 General features
Deg/HtrA proteases belong to a widely conserved enzyme family found across nearly all species. They act independently of ATP and are involved in a variety of important processes. In humans, they are for instance involved in cell signalling, apoptosis, ageing processes and diseases like arthritis and Parkinson’s disease [Strauss, K et al. 2005; Yang, Z et al. 2006]. Higher plant and prokaryotic Deg/HtrA proteases participate in tolerance to stressful conditions and counteract protein folding stress. Plant Deg/HtrA proteases are furthermore involved in degradation of PSII. Some Deg/HtrA proteases, like E.
coli DegP and Saccharomyces cerevisiae (S. cerevisiae) Nma111p (nuclear mediator of apoptosis), can fulfil both protease and chaperone functions depending on temperature [Padmanabhan, N et al. 2009;
Spiess, C et al. 1999]. In contrast, E. coli DegS is a highly specialised enzyme with one known substrate and no detectable chaperone function [Alba, B et al. 2001; Sun, X et al. 2010a].
The name “Deg” is derived from E. coli DegP, the first discovered member of this family. Mutants lacking DegP were unable to degrade periplasmic proteins under stress conditions (DegP for “Degradation of periplasmic proteins”; [Strauch, K et al. 1988]). The same protein was identified in a screen for reduced growth ability at high temperatures, thus leading to the name “HtrA” for “high temperature requirement A” [Lipinska, B et al. 1989]. Deg/HtrA proteases are classified into the MEROPS clan PA and further into the S1 subfamily. That means the active site in the protease domain always includes the triad histidine-aspartate-serine with the latter mediating catalysis. Protease cleavage sites in a substrate are labelled starting from the cleavage site in ascending numbers marked with a prime towards the C- terminus and in ascending numbers towards the N-terminus (Figure 4). Cleavage specificity in the Deg/HtrA family is defined by the amino acid at the P1 position, which is always a hydrophobic residue for Deg/HtrA proteases. Therefore, they are enzymes with chymotrypsin-like specificity.
Figure 4: Cleavage site specificity in substrates
Amino acid residues in the substrate that undergo cleavage are named P1 to P4 in the N-terminal direction and P1’
to P4’ in the C-terminal direction. The substrate specificity pocket residues are named S1 to S4 towards the N- terminus and S1’ to S4’ towards the C-terminus. Protease and proteinase are synonymous.
(Nomenclature was adopted from [Schechter, I et al. 1967; Schechter, I et al. 1968], picture taken from http://web.expasy.org/peptide_cutter/peptidecutter_enzymes.html).
All structurally characterised Deg/HtrA proteases form homotrimers as the basic unit and share common features in domain architecture (Figure 5). The N-terminus contains a transit peptide, a transmembrane anchor or additional domains like the insulin growth factor binding domain (IGFBP) and the Kazal protease inhibitor (KI) domain, of which the structure and function remains to be clarified. At the C-terminus of Deg/HtrA proteases, zero to four PDZ domains are present.
PDZ domains were named after the three Drosophila melanogaster (D. melanogaster) proteins for which this structural element was first described: PSD-95 (postsynaptic density protein), Disc large protein, and zonula occludens protein 1 [Ponting, C et al. 1995]. PDZ domains are involved in substrate recognition and binding and, in case of the Deg/HtrA proteases, regulation of protease activity and oligomerisation (reviewed in [Hansen, G et al. 2013]). PDZ domains are short 80-100 amino acid sequences that mediate specific interactions between proteins by binding to the last five to seven C-terminal amino acid residues of the substrate, which enables them to recognise different substrates [Sheng, M et al. 2001].
Figure 5: Domain architecture of selected Deg/HtrA proteases
All members of the Deg/HtrA family contain a trypsin protease domain (green) with the conserved catalytic triad histidine-aspartate-serine. At the C-terminus zero to four PDZ domains are located (yellow). Optional domains at the N-terminus can be signal sequences (squares), transmembrane anchors (vertical lines), IGFBP (blue) or KI sequences (red).
SS: signal peptide; TM: transmembrane anchor; IGFBP: insulin growth factor binding domain; KI: Kazal protease inhibitor domain; E. coli: Escherichia coli; Synechocys: Synechocystis sp. PCC 6803; H. sapiens: Homo sapiens; D.
rerio: Danio rerio; S. cerevis: Saccharomyces cerevisiae; S. pombe: Schizosaccharomyces pombe; C. albicans:
Candida albicans; A. thaliana: Arabidopsis thaliana; D. melano: Drosophila melanogaster. SwissProt/TREMBL accession numbers are E. coli DegS (P31137), E. coli DegP (P09376), E. coli DegQ (P39099), Synechocystis HtrA (P73354), Synechocystis HhoA (P72780), Synechocystis HhoB (P73940), hHtrA1 (Q92743), hHtrA2 (O43464), hHtrA3 (P83110), hHtrA4 (P83105), D. rerio HtrA2 partial sequence (AL773584), S. cerevis. YNM3 (P53920), S. pombe Q9P7S1, C. albicans YNM3-I (sequence from http://www-sequence.stanford.edu/group/candida/), A. thaliana Q8RY22, A. thaliana DegP1 (O22609), D. melano Q9VFJ3 (Picture taken from [Clausen, T et al. 2002]).
The chymotrypsin-like serine protease domain follows the N-terminal structures and is formed by two β- barrels, with the active site located at the interface between two barrels (Figure 6; [Clausen, T et al.
2011; Clausen, T et al. 2002; Krojer, T et al. 2008b]). The interface-forming loops of the N-terminal β- barrel were named LA, LB, and LC; the loops of the C-terminal β-barrel were named L1, L2, and L3 [Perona, J et al. 1995]. The loops LD, L1 and L2 are called the “activation domain”, which receives signals from the PDZ domains and thus changes its structure towards the active state. The “sensor loop” L3 mediates the signal transduction between the PDZ domains and the protease domains [Clausen, T et al.
2011]. Besides the active site, an oxyanion hole and substrate-specificity pockets are included in the protease domain [Clausen, T et al. 2011]. Loop L1 contains the oxyanion hole, which is formed by amino acid residues preceding the catalytic serine residue and stabilises protein cleavage intermediates. Loop L2 contains the substrate-specificity pocket that binds amino acids preceding the P1 cleavage site in substrates.
Access to the active sites is, although reversible, tightly regulated and often facilitated by oligomerisation of the Deg/HtrA proteases (e.g. in E. coli DegP [Hasenbein, S et al. 2007; Krojer, T et al.
2010; Merdanovic, M et al. 2010]). Another regulation mechanism is the binding of an activator peptide to the PDZ domain like in E. coli DegS [Hasselblatt, H et al. 2007; Wilken, C et al. 2004]. An unique activation mechanism was shown for human HtrA1, where loop L3 directly interacts with the substrate and thus facilitates activation of the proteolytic site [Truebestein, L et al. 2011].
Figure 6: Structure of the E. coli DegP protease domain
Illustrated are the structurally important features of the DegP protease domain representative for all Deg/HtrA proteases. The left panel shows a single protease domain of the DegP hexamer, the right panel a single protease domain of the DegP 24 mer. The hexameric state represents the inactive form, where loop LA* protrudes into the active site of the adjacent monomer to interact with loops L1 and L2. This triad blocks the entrance to the catalytically active site and prevents formation of the oxyanion hole and the substrate-specificity pocket. Upon oligomerisation into dodecamers or 24 mers, loop LA* is pulled back, which releases loops L1 and L2 and induces correct formation of the oxyanion hole and substrate-specificity pocket. Blue: Loop LA* (regulatory loop); Pink:
Loop LD (activation loop); Green: Loop L1 (contains the oxyanion hole); Red: Loop L2 (contains the substrate- specificity pocket); Orange: Loop L3 (sensor loop). The stick model shows the catalytic residues histidine 135 and aspartate 105. The catalytically active serine 210 is exchanged for alanine in this model and thus resembles the proteolytically inactive protease form (Picture taken from [Krojer, T et al. 2008b]).
In 2007, Helm and co-workers [Helm, M et al. 2007] analysed the amino acid sequences of the protease domains of known and putative Deg/HtrA proteases for their evolutionary relationship (Figure 7).
Crucial for indexing enzymes into the same group were the overall domain structure and the protease domain sequence rather than the membership to the same species. Group I contains Deg/HtrA proteases that have one (Synechocystis sp. PCC 6803 HhoA) or two (E. coli DegP) PDZ domains at the C- terminus [Jansen, T et al. 2005; Pallen, M et al. 1997]. Despite lacking a functional PDZ domain, plant DEG5 is also included in this group.
Plant Deg/HtrA proteases with a putative PDZ domain and an extended C-terminus are combined in group II, like e.g. A.thaliana DEG10. Groups I and II also contain all Deg/HtrA proteases with a mitochondrial or chloroplast targeting signal. Fungal Deg/HtrA proteases such as S. cerevisiae Nma111p are combined in group III along with A. thaliana and Oryza sativa (O. sativa) DEG7. The general feature of group III proteases is the second, degenerated protease domain located near the C-terminus.
Moreover, the enzymes of this group are twice as long as the proteases of the other groups. Group IV includes Deg/HtrA proteases without a PDZ domain and with a protease domain shifted towards the C- terminus like the plant DEG15 homologues.
