deg/htra proteases of the cyanobacterium
TRANSCRIPT
Deg/HtrA proteases
of the cyanobacterium
Synechocystis sp. PCC 6803
from biochemical characterization to their physiological functions
Lâm Xuân Tâm
Department of Chemistry
Umeå University
901 87 Umeå
Sweden 2015
Copyright©Lâm Xuân Tâm
ISBN: 978-91-7601-223-9
Front cover: "Colonies of the cyanobacterium Synechocystis sp. PCC 6803"
Printed by: KBC, Umeå University
Umeå, Sweden 2015
To my family
v
Table of content
General introduction 1
I. Synechocystis sp. PCC 6803 as a model cyanobacterium 1 I.1. Cyanobacteria 1 I.2. Synechocystis sp. PCC 6803 2
II. Proteases and Protein quality control 4 II.1. Proteases 4 II.2. Protein quality control 5
III. The family of Deg proteases 6 III.1. Diverse biological functions 7 III.2. Conserved structural architecture 7 III.3. Flexible regulation of activity 9
Aim of this thesis 11
The three Synechocystis Deg proteases 12
I. Biological characteristic 12 I.1. Sequence analysis and Tertiary Structures 12 I.2. Oligomerization states 13 I.3. Proteolytic activity 15
II. Intracellular features 16 II.1. Localization 16 II.2. Proteases expression and regulation 18
III. Physiological importance 20 III.1. Impact on metabolic pathways 20 III.2. Impact on transportation/secretion 21 III.3. Impact on cell envelope biosynthesis 22 III.4. Impact on protein quality control network 22 III.5. Impact on cellular motility 23
Overview and Outlook 24
Acknowledgment 26
References 27
vi
Abstract
The family of Deg/HtrA proteases is present in a wide range of organisms
from bacteria, archaea to eukaryota. These ATP-independent serine
endopeptidases play key roles in the cellular protein quality control. The
cyanobacterium Synechocystis sp. PCC 6803, a model organism for studies
on photosynthesis, metabolism and renewable energy, contains three Deg
proteases known as HhoA, HhoB and HtrA. The three proteases are
important for survival in stress conditions, such as high light or temperature.
In my work the biochemical characteristics of each protease were revealed in
vitro and in vivo. In vitro studies performed using recombinant
Synechocystis Deg proteases allowed conclusions about their
oligomerization states, proteolytic activities and tertiary structure. The in
vivo studies addressed their sub-cellular localization, expression and
physiological importance by comparing wild-type Synechocystis cells with
the three single mutants lacking one of the Deg proteases.
HhoA seems to be involved in the cytoplasmic protein quality control. This
protease is regulated post-transcriptional and post-translational:
oligomerization, pH and/or cation-binding are some of the important factors
to stimulate its proteolytic activity. Instead HhoB acts on periplasmic
proteins and seems to be important for the transportation/secretion of
proteins. While it has low proteolytic capacity, it may act as a chaperone. The
stress-induced HtrA functions in the cellular tolerance against
photosynthetic stress; additionally it might act as a protease partner of
HhoB, generating a protease/chaperone complex.
The results presented in this thesis lay the foundation for a better
understanding of the dynamic protein quality control in cyanobacteria,
which is undoubtedly important for various cellular metabolic pathways.
vii
List of paper
This thesis in based on the following papers that will be referred in the text
by the corresponding Roman numerals
I. Huesgen, P. F., Miranda, H., Lam, X. T., Perthold, M., Schuhmann, H., Adamska, I. and Funk, C. (2011)
Recombinant Deg/HtrA proteases from Synechocystis sp. PCC 6803 differ in substrate specificity, biochemical characteristics and mechanism.
Biochem. J. 435, 733–42.
II. Roberts, I. N., Lam, X. T., Miranda, H., Kieselbach, T. and Funk, C. (2012)
Degradation of PsbO by the Deg protease HhoA Is thioredoxin dependent.
PLoS One 7, e45713.
III. Lam, X. T., Aigner, H., Timmerman, E., Gevaert, K. and Funk, C. Proteomic approaches to identify substrates of the three Deg/HtrA
proteases of the cyanobacterium Synechocystis sp. PCC 6803. Submitted
IV. Hall, M., Lam, X. T., Funk, C. and Persson, K. Structure of the HhoA protease from Synechocystis sp. PCC 6803. Manuscript
viii
Abbreviations
2D-DIGE Two-dimensional difference gel electrophoresis
ADP/ATP Adenosine diphosphate / Adenosine triphosphate
CBB cycle Calvin-Benson-Bassham cycle
COFRADIC Combined fractional diagonal chromatography
GAP Glyceraldehyde 3-phosphate
NADP+/NADPH Nicotinamide adenine dinucleotide phosphate
PSI Photosystem I
PSII Photosystem II
Syn_Degs Synechocystis sp. PCC 6803 Deg proteases
WT Wild-type
GENERAL INTRODUCTION
1
General introduction
I. Synechocystis sp. PCC 6803 as a model
cyanobacterium
I.1. Cyanobacteria, commonly known as blue-green algae, are a
diverse group of photosynthetic prokaryotes that can be found in a wide
variety of habitats. Their name derives from their appearance due to their
green photosynthetic pigment chlorophyll a and the blue phycobilin pigment
phycocyanin. Cyanobacteria are evolutionary the first organisms with a
photosynthetic apparatus containing two photosystems (PS I and PS II)
connected via the cytochrome b6f complex [1] (Figure 1).
Figure 1 - General mechanism of photosynthesis. The absorption of light energy (light
reaction) and the assimilation of CO2 (dark reaction) take place in two separated compartments.
In the light reaction, photons are absorbed by pigments associated with the photosystems (PS I
and PS II). The photosystems as well as the cytochrome b6f complex are membrane-bound
protein complexes located in a specialised membrane system called the thylakoid. Absorbed
photons are funnelled to the reaction centre and trigger an electron flow within and between the
two photosystems. Water is oxidized within PS II producing O2, while NADP+ is reduced to
NADPH within PS I. Water oxidation and electron transport create a proton gradient across the
thylakoid membrane that powers the production of ATP by the enzyme ATP synthase. In the
dark reaction, also known as CBB cycle, ATP and NADPH provide energy for carbon fixation
producing glyceraldehyde 3-phosphate, which can be used for the biosynthesis of sugars, starch
and other organic compounds.
2
Electron transport via two photosystems generates a redox potential strong
enough to use water as an electron donor. The oxygen produced as a
byproduct accumulated and changed the Earth's atmosphere, providing the
basis for evolution of aerobic respiration and with it of almost all live forms
on Earth today. The chloroplast of eukaryotic photosynthetic organisms
(higher plants and algae) has evolved from free living cyanobacteria by
endosymbiosis [2]. Studies on cyanobacteria, therefore, allow insight into an
ancient form of photosynthesis and at the same time provide information for
agriculture and forestry.
Recently, interest in cyanobacteria for their great potential as cell factories
has increased tremendously. Many scientists believe that this is a promising
approach to solve some of the most urgent problems humanity is facing in
the 21th century. Fossil fuels are not just causing problems like pollution and
global warming, these non-renewable sources of energy are also limited in
amount. Conversion of agricultural crops into bio-fuels is less efficient as this
process directly competes to food production. Cyanobacteria, on the other
hands, have high growth rate and can be grown on non-arable land. Even
though cyanobacteria do not naturally produce bio-fuels in usable amounts,
they are easily accessible for genetic engineering. Genetically modified
cyanobacteria were shown to be promising candidates for the production of
bio-fuels and other high value compounds like 1-butanol, ethanol, ethylene,
isobutylaldehyde, isoprene, lactic acid, etc. [3][4][5][6][7]. Thorough
understanding of cyanobacterial proteomes and metabolism, however, is
crucial for further efficient genetic engineering.
I.2. Synechocystis sp. PCC 6803, hereafter: Synechocystis, is the
first cyanobacterium and also the first photosynthetic organism, of which the
entire genome has been completely sequenced [8]. Its known genome,
together with its natural competence and the ability to take up foreign DNA
from the environment, render Synechocystis an excellent model organism
for scientific research. Accordingly, molecular biological modifications
within the genome of Synechocystis, such as inactivation of a specific gene,
can be easily achieved, because the bacterium performs homologous
recombination [9].
Even though Synechocystis was first isolated from a fresh water lake, it also
survives in marine media [10]. Under laboratory maintenance, several sub-
strains of Synechocystis have been derived from the first isolated one [11].
GENERAL INTRODUCTION
3
Glucose-tolerance was bestowed on this organism by unidentified mutations
[9]. In addition to photoautotrophic growth, the glucose-tolerance strain can
grow under mixotrophic, photoheterotrophic or heterotrophic conditions
with glucose as a carbon source [12][13]. The Kazusa strain, of which the
entire genome has been sequenced, is glucose-tolerant.
Figure 2 - Model of a Synechocystis cell with distinct membrane layers and
separated compartments. Pairs of thylakoid membranes enclosing the thylakoid lumen
form several sheet layers and occupy the outer space of the cytoplasm. These membranes
converge at several sites adjacent to the plasma membrane. Carboxysomes usually located in the
central cytoplasm are protein-enclosed micro-compartments containing enzymes of the CBB
cycle.
Synechocystis cells are globular ( 1.5 µm in diameter) and pose the typical
structure of gram-negative cyanobacteria (Figure 2). The outer border
consists of a cytoplasmic membrane (plasma membrane), a mesh-like
peptidoglycan layer, an outer membrane and a lattice surface layer (S-layer)
[14]. Synechocystis also has surface pili supporting twitching motility [15]
and, therefore, is able to move towards a light source (phototactic
movement) [16]. Like all prokaryotes, Synechocystis does not contain a
nucleus or other membrane bound organelles. In order to perform
photosynthesis, Synechocystis has a system of thylakoid membranes that
contain the photosystems and the respiratory complexes. Biogenesis of the
thylakoid membrane in cyanobacteria is not yet understood, they might have
derived from the plasma membrane [17]. However, electron microscopic
examination of Synechocystis cells showed no evidence of association
between the two membranes [18][14].
4
II. Proteases and Protein quality control
II.1. Proteases or peptidases are proteolytic enzymes hydrolyzing
covalent peptide bonds between amino acids of proteins. Exo-proteases act
on the terminal peptide bonds (amino-proteases or carboxy-proteases),
while endo-proteases cleave proteins internally. The general mechanism of
proteolysis involves a nucleophilic attack on the peptide carboxyl group.
Depending on the nucleophile, proteases are traditionally classified into six
groups: Serine, Cysteine, Threonine, Aspartic acid, Glutamic acid and
Metallo proteases. Serine, cysteine and threonine proteases use a functional
group within the enzyme itself, while aspartic acid, glutamic acid and metallo
proteases activate a water molecule to serve as a nucleophile (Figure 3).
Modern classification (MEROPS database [19]) divides proteases into
specific clans (super-families), families and subfamilies according to their
evolutionary origin. Proteases of the same super-family share the same
tertiary structure, but can show divergent evolution to different nucleophiles.
Figure 3 - Proteolytic mechanisms catalysed by different protease groups.
Proteases can be highly specific and only cleave the peptide bond between
certain amino acids. This normally results in limited proteolysis, creating
two or several polypeptides. Limited proteolysis is important in cellular
regulation involving activation or deactivation of certain proteins, removal of
signal peptides, etc. Conversely, other proteases perform unlimited
GENERAL INTRODUCTION
5
proteolysis and totally degrade a protein. Unlimited proteolysis is necessary
to remove malfunctional, misfolded or unwanted proteins and to release
amino acids for new protein synthesis. Proteases performing this
"housekeeping" task play a key role in the cellular protein quality control.
II.2. Protein quality control is a mechanism maintaining
functional proteins for a vast array of cellular activities. Proteins have limited
lifetime, constant synthesis and degradation of proteins are therefore crucial
for cell viability. Newly synthesized polypeptides need to pass
unfolded/partially-folded states before gaining their native three-
dimensional structures and, in many cases, being secreted into specific
intracellular compartments or assembled into specific oligomeric protein
complexes. Under certain (e.g. stress) conditions native proteins can lose
their structures and become misfolded. These non-native protein structures
are kinetically unstable and an efficient network of molecular chaperones
and proteases has evolved to prevent their aggregation [20] (Figure 4).
Figure 4 - Chaperone and protease activities in protein quality control. Transition
between different folding states can occur naturally (gray arrows) or with the support of
chaperones/proteases (dark arrows) (Adapted from [21])
Chaperones are a diverse group of proteins that assist the non-covalent
folding or unfolding of proteins and the assembly or disassembly of protein
complexes without becoming part of the final structure [22]. Chaperones,
therefore, can either assist refolding of damaged proteins or their total
unfolding for efficient degradation. In protein quality control, proteolysis
6
and chaperone activities are often closely connected. Proteases often contain
special subunits or domains for chaperone activity. The proteolytic core
(ClpP) of the Clp protease associates with particular chaperone subunits
(ClpA or ClpX) [23], while FtsH proteases contain AAA domains that unfold
the substrates [24]. Some Deg proteases have been shown to contain dual
protease-chaperone functions [25][26][27].
III. The family of Deg proteases
Deg proteases, formerly DegP (degradation of periplasmic proteins) or HtrA
(high temperature requirement A), are a subfamily of homologous proteases
within the family of chymotrypsin-like serine endopeptidases (S1) according
to current MEROPS classification [19]. In general the structure of Deg
proteases consist of an N-terminal proteolytic domain containing the His–
Asp–Ser catalytic triad followed by one or two C-terminal PDZ domains
mediating a wide range of protein–protein interactions. Unlike other
protease machineries (ClpAP, ClpXP, Lon, FtsH, etc.), Deg proteases contain
no ATP-binding domain and therefore function in an ATP-independent
manner [28].
Figure 5 - Phylogenetic tree of some Deg protease members from A. thaliana
(Ath), E. coli (Eco), human (Hsa) and Synechocystis (Syn). Shown is a simplified
version of a phylogenetic tree constructed around the gene encoding Synechocystis HhoB
(sll1427) by PhylomedDB [29].
GENERAL INTRODUCTION
7
III.1. Diverse biological functions
Deg protease members were first described in E. coli [30][31] and later
detected in other bacteria, archaea and eukaryota. They are involved in
various cellular pathways related to stress tolerance or protein folding during
stress [32]. Some Deg proteases of bacteria (E. coli), mammals (human) and
higher plants (A. thaliana) have been carefully characterized. The E. coli
genome encodes 3 Deg proteases known as DegP, DegQ and DegS (Figure 5).
Periplasmic DegP of E. coli, functioning as a chaperone at normal
temperature and as a protease at high temperature [25], is important for
protein folding and degradation in the cell envelope. The membrane bound
DegS is a regulating protease that can sense misfolded proteins and release
the envelope-stress response signal σE by degrading its inhibition factor [33].
The physiological function of E. coli DegQ is not yet known, however, its
activity appears to be mediated by pH [34]. In humans there are 4 Deg
protease orthologues (HtrA1, HtrA2, HtrA3 and HtrA4); of those best
characterized are HtrA1, acting as tumour suppressor [35], and HtrA2,
involved in mitochondrial protein quality control and regulation of
programmed cell death [36].
In A. thaliana, there are in total sixteen Deg protease members detected in
various organelles (the nucleus, the chloroplast, the mitochondria and the
peroxisome) [37]. Some A. thaliana Deg proteases can have multiple-
localization and at least seven proteases are present in the plastid. So far,
best characterized are the lumenal Deg proteases (Deg1, Deg5 and Deg8)
involved in maintenance of PS II; Deg1 was shown to support the assembly
of Photosystem II [38], while Deg5 and Deg8 can form hetero-complexes
and have synergistic functions in the primary cleavage of the PSII reaction
center protein D1 [39]. These data raise questions concerning the general
role of Deg proteases in photosynthetic organisms [40].
III.2. Conserved structural architecture
Tertiary structures have been completely or partially determined of Deg
proteases in E. coli (DegP, DegQ, DegS) [41][34][33], human (HtrA1, HtrA2,
Htra3) [42][43][44], A. thaliana (Deg1, Deg2, Deg5, Deg8) [45][26][46], T.
maritima (HtrA) [27], L. fallonii (DegQ) [47] and S. pneumoniae (HtrA)
[48]. The protease domains are highly conserved in these organisms
displaying a PA clan (MEROPS classification) with two β-barrels
8
asymmetrically arranged beside each other bringing together the catalytic
triad (His and Asp on the N-terminal barrel and the nucleophilic Ser on the
C-terminal one) [49][50] (Figure 6). The small PDZ domain is less conserved
and consists of two β-sheets with two α-helices on its edges. The two
domains are connected by a flexible linker allowing en bloc mobility and
different arrangements of the PDZ domain.
Figure 6 - Tertiary structure of E. coli DegS, human HtrA2, A. thaliana Deg1 and
Synechocystis HhoA. The protease domains shown in gray (red for HhoA) with two
conserved β-barrels (C-terminal β-barrel in darker colours) containing the regulation loops (LA,
LD, L1, L2 and L3) are arranged in the same angle. The relative positions of the PDZ domains
shown in yellow (orange for HhoA) containing the interaction clamps (IC1 and IC2) compared
to the proteases domain are different in these structures.
GENERAL INTRODUCTION
9
Via hydrophobic interactions between the protease domains Deg proteases
form stable homo-trimeric complexes. PDZ domains and/or the protruding
LA loop of the protease domain facilitate further oligomerisation of these
basic trimer units into bigger complexes such as hexamers, 12-mers or 24-
mers. These cage-like oligomeric structures is suggested to trap substrate
within the inner cavity until the protein processing is complete. Formation of
high-order oligomers can be triggered by the presence of substrate or change
of pH [51] [45] and normally involves an activation mechanism of the
protease.
III.3. Flexible regulation of activity
Under normal condition, Deg proteases are present in a "resting" state which
can reversibly transform to the active state upon specific signals such as the
presence of misfolded polypeptide or ions, changes in pH or temperature,
etc. The activation mechanism can vary considerably in different Deg
proteases (Figure 7).
Activation by formation of higher-order oligomers is reported for E. coli
DegP and A. thaliana Deg1. In the "resting" state, E.coli DegP assembles into
hexamers, in which the active sites are blocked and distorted by the
protruding LA loop of the subunit within the opposite trimer [52]. Substrate-
binding to the PDZ1 domain destabilizes the DegP hexamer leading to
formation of 12- or 24-mers. In the multi-oligomers, the LA loop is
withdrawn from the active site and the PDZ1 domain is fixed to be able to
interact with the L3 loop, which in turn interacts with the loops LD, L1 and
L2 of the adjacent subunit [53]. Consequently, the active site is restructured
into proper conformation for proteolysis. Oligomerization of A. thaliana
Deg1 into the active hexameric state, in which the PDZ domain is fixed for
the PDZ→L3→LD/L1/L2 activation, is triggered by low pH [45].
E. coli DegS, which is normally present in the cell as membrane bound
trimer with disordered active site, is not activated by oligomer conversion.
The PDZ domain of DegS is kept in position by additional interactions with
the protease domain. It captures the L3 loop and therefore prevents
interaction with the loops LD/L1/L2 of the adjacent subunit. Binding of
allosteric activators to the PDZ domain, such as misfolded outer-membrane
proteins, triggers the L3 loop to re-orient and to interact with the loops
LD/L1/L2, and thus, restructures the active site [33].
10
Figure 7 - Schematic demonstration of the activation mechanisms of proteolytic
Deg proteases in E. coli (DegP and DegS) and human (HtrA1 and HtrA2). Trimers of
the Deg proteases are shown with the loops L1, L2, L3, LA and LD important for activity
regulation.
Different to E. coli Deg proteases, in human HtrA1 the PDZ domain is not
involved in the activation mechanism, its removal therefore has no effect on
the proteolytic activity of HtrA1 [42]. The disordered active site of the HtrA1
trimer can be oriented and stabilized by substrate binding via an induced-fit
mechanism. Binding of polypeptides to the active site and the L3 loop is
sufficient to activate HtrA1. To regulate activity of HtrA1 therefore likely
requires additional control mechanisms.
Finally, the membrane bound human HtrA2 trimer maintains its properly
oriented active site in the "resting" state. Access to the active site, however, is
restricted by the PDZ domain [44]. Certain conditions (e.g. substrate
binding) can induce relocation of the PDZ domain and allow proteolysis.
HtrA2 depleted of the PDZ domain was shown to have high proteolytic
activity.
AIM OF THIS THESIS
11
Aim of this thesis
Three members of the Deg protease family have been detected in
Synechocystis (Syn_Degs), known as HhoA (sll1679), HhoB (sll1427) and
HtrA (slr1204), which protect Synechocystis against high light or heat stress
[54]. Little is known about their course of action (e.g. their substrates,
regulation mechanisms, etc.).
My studies aimed for a more detailed view on the cyanobacterial Deg
proteases. In vitro and in vivo analyses were performed on recombinant
truncated Syn_Degs (rHhoA, rHhoB and rHtrA) and deletion mutants with
each of the three deg genes inactivated (ΔhhoA, ΔhhoB and ΔhtrA) (Figure
8). The results highlighted the function of each protease, which showed to be
individual for each protease.
Figure 8 - Summary of scientific research on Synechocystis Deg proteases carried
out in this thesis. Soluble recombinant Syn_Degs were constructed as His-tagged, truncated
versions depleted of the predicted signal peptide (for rHhoA and rHhoB) or transmembrane
domain (for rHtrA) (PAPER I). Additionally stable inactive versions were constructed of the
recombinant Deg proteases, in which the catalytic Ser residue was mutated to Ala. Single deg
mutants (ΔhhoA, ΔhhoB and ΔhtrA) were generated by replacing a fragment in the middle of
each individual deg gene (hhoA, hhoB or htrA) with a kanamycin-resistance marker, thus,
preventing the expression of a functional protease (PAPER III).
12
The three Synechocystis Deg proteases
I. Biological characteristic
I.1. Sequence analysis and Tertiary Structures
The three Synechocystis Deg proteases share around 50 % mutual sequence
identity and are evolutionary more closely related to each other than to Deg
proteases found in other organisms [55].
Figure 9 - Multiple sequence alignment of the protease domain and the first PDZ
domain belonging to Deg proteases of Synechocystis (HhoA, HhoB and HtrA), E.
coli (DegP, DegQ and DegS), human (HtrA1 and HtrA2) and A. thaliana (Deg1,
Deg5 and Deg8). Positions of the catalytic triads (His–Asp–Ser) and the regulation loops (LA,
LD, L1, L2 and L3) on the protease domain as well as the carboxylate-binding loop and the
interaction clamps (IC1 and IC2) on the PDZ domain are marked above the sequences.
Alignment were generated using Clustal Omega [56] and coloured by conservation.
THE THREE SYNECHOCYSTIS DEG PROTEASES
13
The protease domains of Syn_Degs are highly conserved to other Deg
proteases, while the PDZ domains are variable. The tertiary structure of
hexameric HhoA was solved by X-ray crystallography (PAPER IV) (Figure 6).
In this "resting" state, the loops L1, L2 and L3 are disordered, thus, a
functional catalytic triad, oxyanion hole and S1 binding pocket are lacking.
Furthermore, no electron density of the LA loop could be observed indicating
that this loop is disordered in the current HhoA structure and not required
for hexamer formation. All three Syn_Degs contain relatively long LA loop
and IC1 compared to other Deg proteases (Figure 9). Dimerization of HhoA
trimers relies solely on the PDZ domains; IC1 of a subunit within trimer 1
interacts with IC2 of the opposite subunit belonging to trimer 2.
In the PDZ domain, the carboxylate-binding loop, which is required for
recognition of substrate C-termini, has a YIGV motif in HhoA and a YLGI
motif in HhoB or HtrA. Interestingly, the Arg residue placed in front of the
carboxylate-binding loop, which is highly conserved in Deg proteases of
other organisms, is exchanged for a His in all three Syn_Degs (Figure 9). In
resolved structure of HhoA, this His286 residue is rotated and pointing away
from the carboxylate-binding pocket. By forming a salt bridge with the
conserved Asp154 of the protease domain, it determines the relative
orientation of the two domains and accordingly could affect the formation of
higher-order oligomers or PDZ–L3 interaction. In E. coli DegS, an
equivalent salt bridge between Arg and Asp residues is important for the
PDZ→L3 activation [33]. It should be noted that the proteolytic activity of
the three Syn_Degs is very low at pH ≤ 5,0 (PAPER I), at conditions favoring
a charged histidine side chain and salt bridge formation with Asp.
A Mg2+ hexa-hydrate ion was observed to be bound by the conserved Asp229
residue of the L1 loop at the central channel of the HhoA trimer unit. Binding
of Ca2+ to that position is known for the lumenal A. thaliana Deg5 [46].
Binding of divalent cations may regulate protease activity allosteric as the
proteolytic activity of the three Syn_Degs highly increased in the presence of
Ca2+ or Mg2+ (PAPER I).
I.2. Oligomerization states
The oligomerization states of recombinant Syn_Degs with different
molecular weights were determined by gel filtration (PAPER I). Among the
three recombinant proteases, rHhoA had the highest ability to form
14
oligomers, it was present hexameric, whereas rHhoB and rHtrA only formed
trimers. The presence of the substrate β-casein induced the formation of
large oligomers, rHhoA formed 12-24-mers, rHtrA 9-12-mers, however,
rHhoB remained as trimer.
It should be noted that native HtrA is expected to anchor into the cellular
membrane, and therefore aggregation into larger homo-oligomers might be
prevented in vivo. Based on immuno stains (PAPER III) its transmembrane
domain, which has been omitted for construction of the recombinant
protease (PAPER I), seems to be intact in vivo. It therefore is unlikely that
HtrA is released from the membrane by N-terminal cleavage and may
function as trimer or hetero-oligomer. Other membrane-bound Deg
proteases, like E. coli DegS or human HtrA2, have been shown to function as
trimer [36][57].
Figure 10 - Structure of the HhoA hexamer. Trimer formation is mediated by the
proteases domains (cyan), while dimerization of the two trimers relies on the PDZ domains
(blue). (Adapted from PAPER IV).
Hexameric rHhoA was analysed by X-ray crystallography (PAPER IV). The
cage-like structure forms three entrance pores allowing only small or
unfolded polypeptides to access the catalytic inner cavity (Figure 10). The
PDZ domains form three pillars within the hexameric structure and are
responsible for trimer interaction. Accordingly, rHhoA, and also rHtrA,
depleted of the PDZ domains only will be present in trimeric state and lose
the ability to form larger oligomers, even in the presence of substrate
(PAPER I).
THE THREE SYNECHOCYSTIS DEG PROTEASES
15
I.3. Proteolytic activity
The proteolytic activity of recombinant Syn_Degs were examined against
different model substrates (PAPER I) or the whole Synechocystis proteome
(PAPER III). In general, rHhoA was shown to be the most active among the
three proteases, while rHhoB was almost proteolytic inactive (Figure 11).
Deletion of the PDZ domain, strongly impaired, but did not abolish the
proteolytic activity of rHhoA or rHtrA.
Figure 11 - In vitro activity of recombinant Synechocystis Deg proteases against
different purified proteins. All three proteases can process naturally unfolded β-casein, but
not natively folded BSA, lysozyme or PsbO from either Synechocystis or the higher plant
spinach. Reduced lysozyme or PsbO, however, could be degraded significantly by rHhoA.
(Adapted from PAPER I and PAPER II)
Using model substrates, the proteolytic activity of the three proteases
differed between the three proteases and was dependent on pH, temperature
(Figure 12) or the presence of Ca2+/Mg2+. rHhoA was highly active over a
broad pH (from pH 5 to pH 8) and temperature range. Its activity increased
tremendously with raised temperature up to 55oC, then dropped rapidly
(probably due to protease denaturation). rHhoB, on the other hand, showed
no significant proteolytic activity. Instead chaperone activity was detected
for rHhoB, but not for rHhoA or rHtrA using the refolding assay of
denatured E. coli α-amylase MalS (PAPER III). rHtrA was proteolytic active
at relative acidic pH (pH 5 - 6,5) and within a narrow range of temperature.
In contrast to rHhoA, its activity was highest at the optimal growth
temperature of Synechocystis (33 oC) [58] and decreased slowly with raised
temperature.
16
Figure 12 - pH/Temperature profile of recombinant Synechocystis Deg proteases
activity without Ca2+/Mg2+. The proteolytic activity of the three proteases was examined
using casein substrate labelled with pH-insensitive fluorescent dyes. While the fluorescent
signal is quenched in intact substrate, it will be released during degradation into smaller
fragments. (Adapted from PAPER I)
Addition of Ca2+ or Mg2+ activated rHhoB, and strongly enhanced the
proteolytic activity of rHhoA and rHtrA. The common effect of divalent ions
on the three proteases suggests a general activation mechanism. Ca2+/Mg2+-
binding may trigger structural changes of either the proteases or the
substrates and favour proteolysis. Binding of Ca2+ or Mg2+ has been reported
for rHhoA (PAPER IV) as well as for A. thaliana Deg5 [46]. In the presence
of Ca2+/Mg2+, the pH profiles of recombinant Syn_Degs was broadened
towards basic pH (PAPER I).
In total Synechocytis protein extracts, recombinant Syn_Degs tend to
degrade the same substrates, but prefer different cleavage sites (PAPER III).
rHhoA and rHtrA also were shown to degrade the model substrate β-casein
at different sites (PAPER I). All three Syn_Degs preferred hydrophobic
residues in positions P4 to P1'. High preference for small amino acids like
Val or Al were observed at P1 and Ala or Ser at P1' positions indicating small
or narrow substrate binding pockets at the active sites of the three proteases.
II. Intracellular features
II.1. Localization
Syn_Degs have been detected in multiple subcellular compartments of
Synechocystis isolated by combined method including gradient
THE THREE SYNECHOCYSTIS DEG PROTEASES
17
centrifugation and two-phase partitioning. HhoA was detected as soluble
protein in the periplasm [59] and also seems to be bound to the plasma
membrane and thylakoid membrane [60] (PAPER II). HhoB was only found
to be associated with the plasma membrane, while HtrA was detected in
outer membrane [61], inner membrane and thylakoid membrane fractions
(PAPER II). These biochemical data, however, do not reveal, whether the
Deg proteases are located on the periplasmic, cytoplasmic or lumenal side of
the membrane.
Specific polyclonal antisera directed against recombinant Syn_Degs allowed
the localization of HhoA in the cyanobacterial cell using gold immuno-
labelling. HhoA was predominantly detected on the cytoplasmic side of the
thylakoid membranes and also concentrated on the septum of dividing cells
(Figure 13). This result is consistent to the proteomic detection of HhoA in
various samples and further shows the cellular ability to distribute HhoA to
different compartments. Apparently, the presence of HhoA in the periplasm
is only important during the division stage of non-stressed cells. Septal
localization of Deg proteases has also been reported for S. pneumoniae [62].
Figure 13 - Immuno-gold labelling of HhoA in the triple mutant depleted of all
three Deg proteases (Δ3), wild-type (WT) and the ΔhhoB-htrA double mutant (Δ2)
[54] (Bar = 0,5 µm). HhoA molecules were visualized by gold particles and appeared as small
dark spots. The immuno response to HhoA was much stronger in Δ2 mutant than in WT
suggesting an up-regulation of HhoA in the absence of other Deg proteases.
18
II.2. Proteases expression and regulation
The protein expression of Syn_Degs in WT and single deg mutants grown at
normal conditions or after exposure to high light or heat stress were
examined by immunoblotting using inactive recombinant Syn_Degs as
quantitative standards (PAPER III). The three Syn_Degs showed distinct
regulation under light/heat stress treatment. Their expression was strongly
affected by the absence of other Deg proteases, especially HhoB and HtrA
seemed to be regulated inversely correlated (Figure 14).
Figure 14 - Protein expression of
Synechocystis Deg proteases in
wild-type and the three single deg
mutants grown at normal conditions
and in response to light or heat
stress. The two forms of HhoA with
higher and lower molecular weight are
presented as upper and lower bars of the
stacked columns. (PAPER III)
Figure 15 - Activity-based profiling of
all serine hydrolases in
Synechocystis. Labelling reaction with
biotinylated fluorophosphonate was
carried out on wild-type and single deg
mutants either on total extract (TE),
loaded on equal Chl a concentration, or
the soluble fraction (Sol), isolated from the
same amount of wild-type TE. The
expected Syn_Degs positions are marked
with arrowheads (red for HhoA, green for
HhoB and blue for HtrA).
The antibody directed against HhoA immuno-stained two protein bands
differing in molecular weight with about 2 kDa, these peptides might be the
result of divergent post-translational modifications in vivo. Expression of
THE THREE SYNECHOCYSTIS DEG PROTEASES
19
HhoA appeared to be stable, a significant increase in the total amount of
HhoA was only observed in ΔhhoB or ΔhtrA cells exposed to light stress.
While there was no obvious change in the total amount of HhoA exposed to
heat stress, the ratio of the two forms changed; the higher molecular weight
band accumulated, while the one with lower molecular weight decreased.
Both these HhoA forms appear to be proteolytic active, as they are labelled
by activity-based profiling (Figure 15); however, only the band with higher
molecular mass was detected to be soluble. In agreement to this finding,
proteomic studies revealed HhoA in the soluble fraction of Synechocystis to
accumulate under heat shock [63].
Furthermore, both forms of HhoA seemed to be activity-labelled 2-3 times
stronger in ΔhhoB and ΔhtrA background than in WT under normal growth
conditions (Figure 15), while there was no obvious difference in the total
amount of that protease (Figure 14). In WT therefore a large portion of HhoA
seems to be proteolytic inactive in a "resting" state. It could be activated
under stress conditions, which would work much faster than up-regulated
transcription of hhoA. It is known that the proteolytic activity of rHhoA
increases strongly at elevated temperature (PAPER I).
The protein amount of HhoB seems to be stable and independent of light
stress, even in the absence of HhoA or HtrA. Under heat stress, however,
HhoB amount decreased significantly, especially in ΔhtrA. Compared to
HhoA or HhoB, HtrA is most stress-induced; its usually low expression
increased considerably in response to high light, heat stress, or to the
absence of HhoB. Gene transcription of htrA also differs to hhoA and hhoB.
After shifting dark adapted cells to light, htrA is transiently expressed; its
mRNA can reach maximal level within 5 min, but then declines to the
amount of normal growth condition, while hhoA transcription is maximal
after 15 min and hhoB transcripts still accumulate slowly after 4 hours [64].
The inverse correlated regulation of HhoB or HtrA in ΔhtrA or ΔhhoB,
respectively, was drastically enhanced during high light or thermal stress,
indicating a connection between their activity. In addition, a similar, but less
obvious negative correlation was observed between HhoA and the
HhoB/HtrA pair (accumulation of HhoA in ΔhhoB/ΔhtrA exposed to light
stress and decrease of HhoB/HtrA in ΔhhoA under heat stress compared to
WT under the same conditions).
20
III. Physiological importance
Differences in the proteomes of single deg mutants compared to WT allowed
conclusions on the biological roles of each protease. The proteomes were
examined based on their differences in expressed proteins (2D-DIGE
method) (Figure 16) and based on naturally occurring N-termini in the
proteome (N-terminal COFRADIC method) (PAPER III). Deletion of the Deg
proteases had impact on various cellular activities; the main metabolic
pathways and the transportation/secretion systems were most strongly
affected.
Figure 16 - Differently regulated proteins in each single deg mutant compared to
Synechocystis wild-type. Proteins, which were detected to accumulate (black letters) in one
mutant were never found to be decreased (white letters) in other mutants and vice versa.
III.1. Impact on metabolic pathways
Deletion of either one of the Syn_Degs lead to changed accumulation of two
important enzymes (Gap2 amount decreased and amount of Pyk2 increased)
controlling the carbon flow from the CBB cycle toward GAP or pyruvate
formation (PAPER III). There are two Gap homologs in Synechocystis; Gap1
only is important in glycolysis and its absence does not affect the
THE THREE SYNECHOCYSTIS DEG PROTEASES
21
photosynthetic growth of Synechocystis, even under light stress; Gap2 on the
other hand is functioning in CBB cycle and gluconeogenesis and is crucial for
photosynthetic growth [65]. Transcription of gap2 also is dependent on the
photosynthetic electron transport and active carbon metabolism [66]. The
decreased amount of Gap2 therefore may indicate impaired photosynthetic
activity in the single deg mutants. The amount of Sll1358, an enzyme
involved in photorespiration, was also diminished in all three mutants.
Interestingly, the amount of Gap2 and Pyk2 varies much stronger from WT
in ΔhhoA than in the other two mutants. Additionally Sll0529, a
transketolase homolog, accumulated only in ΔhhoA, but not the other two
mutants. On the other hand, FbaII, mainly responsible for Fba activity in
Synechocystis [67], and CcmA, essential for carboxysome formation in
Synechocystis [68], were both up-regulated in ΔhhoB or ΔhtrA, but not in
ΔhhoA. Furthermore, deletion of HhoA affected the neo-N-termini of
proteins involved in carbon metabolism (SucD, CP12 and RbcS), while the
neo-N-termini of proteins forming the photosystems or carboxysome were
affected in ΔhhoB (PsbO and PsaC) and ΔhtrA (PsbO, CcmM and PsaD). A
combined in vivo and in vitro COFRADIC approach identified the
cytoplasmic RbcS as a natural substrate for HhoA and the lumenal PsbO as
substrate for both HhoB and HtrA (PAPER III). HhoA may be important for
a functional CBB cycle, while the HhoB/HtrA pair regulates photosynthetic
activity.
III.2. Impact on transportation/secretion
Deletion of single Syn_Degs lead to decreased accumulation of some ABC
transporters (urea precursor UrtA and bicarbonate precursor CmpA) and to
accumulation of neo-N-termini belonging to Sll0141, a HlyD family secretion
protein. Distinct differences in the regulation of the phosphate permease
system was observed only in ΔhhoB, but not the other two mutants (Figure
17). PstS1 obviously was absent in ΔhhoB leading to strong accumulation of
SphX, PstB1' and possibly PstS2 (PAPER III). Furthermore, the COFRADIC
experiment revealed in ΔhhoB regulation of many other transportation
proteins (Sll0180, HlyD, PilJ, Ycf60, Sll7070, Cph1, FutA2, Slr0615, Slr0977
and TonB) based on the accumulation of their natural-N-termini. HhoB
activity therefore might be important for the proper function of several
transportation systems, especially the type I secretion mechanism.
22
Figure 17 - Significant differences of some phosphate import proteins in ΔhhoB
compared to WT detected by 2D-DIGE. PstS1 and SphX are two periplasmic phosphate
precursor homologs, while PstB1' is an ATP-binding subunit of the phosphate permease. (3D
mapping of the 2D-DIGE fluorescent signals generated by SameSpots software).
III.3. Impact on cell envelope biosynthesis
The neo-N-termini of some structural proteins (HlyA and Slr1908) or
enzymes (MurA, MurC and LpxA) involved in cell structure were
significantly differentially regulated upon HhoB deletion (PAPER III).
Changes in HlyA, a Synechocystis S-layer protein [69], and Slr1908
containing porin and S-layer homology domains may be the direct result of
impaired secretion systems in ΔhhoB. MurA and MurC are suggested to
catalyse the cytoplasmic reactions of the peptidoglycan biosynthesis [70],
while LpxA functions in the metabolism of lipopolysaccharides, a major
component of the outer membrane [71].
HtrA deletion lead to the complete disappearance of two neo-N-termini
belonging to the penicillin-binding protein Pbp8. Penicillin-binding proteins
are enzymes catalysing the periplasmic reactions of peptidoglycan
biosynthesis, maturation or recycling [72]. Pbp8, involved in the final step of
Synechocystis cell division [73], was also identified as natural substrate of
HtrA using N-terminal COFRADIC.
III.4. Impact on protein quality control network
Deletion of HhoA lead to accumulation of the neo-N-terminus of the
substrate-binding adaptor ClpS, which can target destabilized N-terminal
residues of substrate polypeptides to the ClpAP/ClpXP proteolytic
THE THREE SYNECHOCYSTIS DEG PROTEASES
23
machinery. The ATP-dependent chaperone ClpC was up-regulated in ΔhhoB
and ΔhtrA. Further on, the natural N-termini belonging to several proteases
(Lons, HhoA, HtrA, Slr1288, TldD and CtpC) strongly accumulated in
ΔhhoB. It is likely that these proteases/chaperones are up-regulated to
compensate for the loss of the specific Deg protease activity in the protein
quality control network.
III.5. Impact on cellular motility
Deletion of the individual Syn_Degs affected several proteins related to
chemotaxis. The type 4 pilin polypeptides PilA1 and PilO decreased in single
deg mutants, in ΔhtrA additionally a neo-N-terminus of the pilus assembly
protein PilQ was diminished (PAPER III). PilA1, PilO and PilQ proteins are
crucial for phototactic motility of Synechocystis [74]. Furthermore, the
amount of Sll0414 was decreased in ΔhhoA and ΔhhoB and Sll0301 had neo-
N-termini accumulating in ΔhhoB. While the function of these two
pentapeptide repeat proteins is still unknown, interruption of either sll0414
or sll0301 will lead to reduced or absent cellular motility [75]. Accordingly,
absence of motility and abnormal pilus assembly in a triple deg mutant
lacking all three Syn_Degs has been observed, but not the double deg
mutants [54].
24
Overview and Outlook
The three Deg proteases of Synechocystis are distinctly different in their
biochemical characteristics. Their suspected physiological functions are
different from each other and seem to be in consistence with previous co-
expression data showing HhoA to be involved in general house-keeping
processes, HhoB co-expressing with periplasmic proteins and HtrA with
electron transport chain proteins [76].
Synechocystis HhoA appears to be a highly active protease functioning over
a wide range of pH and temperature. Unlike HhoB and HtrA, which usually
generate large proteolytic fragments, HhoA can degrade misfolded proteins
completely due to its ability to form higher order oligomers trapping
substrate within its inner cavity. In vivo, in cells growing in normal
condition, a large portion of HhoA is inactivated in a "resting" state. Instead
of being regulated on transcriptional level, several factors such as pH,
temperature influence its activity and with that its molecular weight
(distribution of higher or lower molecular weight form) or even localization.
In non-dividing cells, HhoA seems to be a cytoplasmic protease that mainly
functions on the cytoplasmic site of the thylakoid membrane. However, it
can be found also in the periplasm and in the septum during cell division,
maybe stress conditions induce various locations. Absence of HhoA affects
mainly cytoplasmic proteins, and the cytoplasmic RbcS is proposed as a
natural proteolytic substrate of HhoA. My current results point to the
physiological role of HhoA as a general cytoplasmic "house cleaning"
protease.
Further experiments based on our HhoA crystal structure are needed to
understand the regulation mechanism of its activity. While proteolytic
activation upon binding of substrate polypeptides are well studied in E. coli
Deg proteases, little is known about how other signals, such as pH, ions or
post-translational modifications can regulate proteolysis. Based on the
current HhoA structure, the Ca2+/Mg2+ binding site or the pH-dependent salt
bridge between the protease and PDZ domain might possibly function as
allosteric regulators. Furthermore, the in vivo regulation of the two forms of
HhoA with slightly different molecular weights would be interesting. Such
post-translational modifications have not been observed in other Deg
OVERVIEW AND OUTLOOK
25
proteases. Finally the localization of HhoA at multiple sites within the cell
and even its controlled secretion are interesting topics for further studies.
Synechocystis HhoB has very low proteolytic activity compared to HhoA.
HhoB may be a highly specific protease only degrading certain substrates
under specific conditions, such as E. coli DegS. We cannot exclude the low
activity of recombinant HhoB to be caused by poor construction. Based on
the molecular weight native HhoB seems to contain an intact signal peptide,
which is not present in rHhoB. However, this truncation makes rHhoB more
similar to the highly proteolytic active rHhoA. A more tempting speculation
therefore is that HhoB may function as molecular chaperone. Chaperone
activity was detected for HhoB in vitro. Furthermore, I propose the presence
of HhoB to be crucial, but not sufficient, for the degradation of PsbO in vivo;
additionally the proteolytic active HtrA is required. Absence of HhoB lead to
different expression of several periplasmic proteins, especially the secretion
systems. Depletion of the major phosphate precursor and sensor PstS1 is
unlikely the result of slower protein turnover upon protease deletion. These
changes rather might be triggered by lacking chaperone activity of HhoB,
which is important for protein secretion. My results suggest that while HhoA
or HtrA activity is important under stress conditions, HhoB is required for
normal cellular activity. During stress HhoB activity seems to be tightly
connected to HtrA activity.
Further experiments are needed to verify the hypothetical chaperone activity
of HhoB in vivo. Future studies should explore an involvement of HhoB in
cellular transportation/secretion, in cell envelope biogenesis as well as in
PSII maintenance. The exact localization of HhoB and its co-
regulation/complex formation with HtrA are key questions.
Synechocystis HtrA is a membrane bound protease, with narrow pH- and
temperature-optima. Accordingly, regulation of HtrA activity seems to be
based on rapid expression of the protease. HtrA can degrade Pbp8
independently, but needs HhoB to degrade PsbO. It will be interesting to
investigate if HhoB and HtrA degrade PsbO step-wise or if both proteases
form a hetero-oligomeric complex, similar to A. thaliana Deg5 and Deg8 for
D1 degradation. Further experiment are needed to examine the
oligomerization states and their influence on proteolytic activity of a
HhoB/HtrA mixture.
26
Acknowledgment
My doctoral studies have received financial support from the Swedish
Energy Agency, the Umeå University, the European Union 7th Framework
Program (through the Prime-XS proteomic project) and the Lawsky
foundation.
First of all, I would like to thank my supervisor, Prof. Christiane Funk,
who has given me opportunities, conditions, guidance and a lot of
encouragements for my professional development. Thank you for your
understanding, patience and generosity, Doktormutter!
I would also like to thank all my colleagues; those who have involved in my
work directly (Harald Aigner, Helder Miranda, Irma Roberts, Pitter
Huesgen, Michael Hall, Evy Timmerman, Prof. Kris Gevaert,
Samantha Bryan and Prof. Conrad Mullineaux) and some who have
given me valuable instructions, advices and opinions (Otilia Cheregi,
Tania Tibiletti, Giada Marino, Prof. Wolfgang Schröder, Patrick
Storm, Miguel Hernandez, Raik Wagner, Marcus Wallgren, Assoc
Prof. Anders Hofer, Aaron Edwin, Prof. Uwe Sauer and many others
that I don't have the ability to list here).
Lastly but above all, I want to show my deepest gratitude to my family
especially my wonderful parents (Lâm Văn Thuận and Phạm Thị Xuân),
who have devoted their life to my development in every possible ways and
my husband Harald Aigner, who has walked into my life at the beginning
of my doctoral studies and ever since then, support me and keep me from
being lost both in my professional and personal life. Thank you for always
believing in me more than myself and for maintaining my belief in good
people and good scientists!
Special thanks to my friends
in Umeå especially the
delightful Vietnamese
community and the late
beloved Lars Lundmark.
You all have been a warm
part of my Swedish life!
REFERENCES
27
References
[1] R. E. Blankenship, “Origin and early evolution of photosynthesis,”
Photosynth. Res., vol. 33, no. 2, pp. 91–111, 1992.
[2] J. A. Raven and J. F. Allen, “Genomics and chloroplast evolution: what
did cyanobacteria do for plants?,” Genome Biol., vol. 4, no. 3, p. 209,
2003.
[3] E. I. Lan and J. C. Liao, “ATP drives direct photosynthetic production
of 1-butanol in cyanobacteria,” Proc. Natl. Acad. Sci. U. S. A., vol. 109,
no. 16, pp. 6018–23, 2012.
[4] M. D. Deng and J. R. Coleman, “Ethanol synthesis by genetic
engineering in cyanobacteria,” Appl. Environ. Microbiol., vol. 65, no.
2, pp. 523–8, 1999.
[5] K. Takahama, M. Matsuoka, K. Nagahama, and T. Ogawa,
“Construction and analysis of a recombinant cyanobacterium
expressing a chromosomally inserted gene for an ethylene-forming
enzyme at the psbAI locus,” J. Biosci. Bioeng., vol. 95, no. 3, pp. 302–
5, 2003.
[6] S. Atsumi, W. Higashide, and J. C. Liao, “Direct photosynthetic
recycling of carbon dioxide to isobutyraldehyde,” Nat. Biotechnol., vol.
27, no. 12, pp. 1177–80, 2009.
[7] S. A. Angermayr, M. Paszota, and K. J. Hellingwerf, “Engineering a
cyanobacterial cell factory for production of lactic acid,” Appl.
Environ. Microbiol., vol. 78, no. 19, pp. 7098–106, 2012.
[8] T. Kaneko, S. Sato, H. Kotani, a. Tanaka, E. Asamizu, Y. Nakamura, N.
Miyajima, M. Hirosawa, M. Sugiura, S. Sasamoto, T. Kimura, T.
Hosouchi, a. Matsuno, a. Muraki, N. Nakazaki, K. Naruo, S. Okumura,
S. Shimpo, C. Takeuchi, T. Wada, a. Watanabe, M. Yamada, M.
Yasuda, and S. Tabata, “Sequence Analysis of the Genome of the
Unicellular Cyanobacterium Synechocystis sp. Strain PCC6803. II.
Sequence Determination of the Entire Genome and Assignment of
Potential Protein-coding Regions,” DNA Res., vol. 3, no. 3, pp. 109–
136, 1996.
28
[9] J. G. K. Williams, “Construction of specific mutations in photosystem
II photosynthetic reaction center by genetic engineering methods in
Synechocystis 6803,” Methods Enzymol., vol. 167, pp. 766–778, 1988.
[10] D. L. Richardson, R. H. Reed, and W. D. P. Stewart, “Synechocystis
PCC6803: a euryhaline cyanobacterium,” FEMS Microbiol. Lett., vol.
18, no. 1–2, pp. 99–102, 1983.
[11] M. Ikeuchi and S. Tabata, “Synechocystis sp. PCC 6803 - a useful tool
in the study of the genetics of cyanobacteria,” Photosynth. Res., vol.
70, no. 1, pp. 73–83, 2001.
[12] T. Nakajima, S. Kajihata, K. Yoshikawa, F. Matsuda, C. Furusawa, T.
Hirasawa, and H. Shimizu, “Integrated metabolic flux and omics
analysis of Synechocystis sp. PCC 6803 under mixotrophic and
photoheterotrophic conditions,” Plant Cell Physiol., vol. 55, no. 9, pp.
1605–12, 2014.
[13] S. L. Anderson and L. Mcintosh, “Light-activated heterotrophic
growth of the cyanobacterium Synechocystis sp . strain PCC 6803: a
Blue-Light-Requiring Process,” J. Bacteriol., vol. 173, no. 9, p. 2761,
1991.
[14] A. M. L. van de Meene, M. F. Hohmann-Marriott, W. F. J. Vermaas,
and R. W. Roberson, “The three-dimensional structure of the
cyanobacterium Synechocystis sp. PCC 6803,” Arch. Microbiol., vol.
184, no. 5, pp. 259–70, 2006.
[15] D. Bhaya, N. R. Bianco, and D. Bryant, “Type IV pilus biogenesis and
motility in the cyanobacterium Synechocystis sp . PCC6803,” Mol.
Microbiol., vol. 37, no. 4, pp. 941–951, 2000.
[16] D. Bhaya, a Takahashi, and a R. Grossman, “Light regulation of type
IV pilus-dependent motility by chemosensor-like elements in
Synechocystis PCC6803,” Proc. Natl. Acad. Sci. U. S. A., vol. 98, no.
13, pp. 7540–5, 2001.
[17] C. Mullineaux, “The thylakoid membranes of cyanobacteria: structure,
dynamics and function,” Aust. J. Plant Physiol., vol. 26, no. 7, p. 671,
1999.
REFERENCES
29
[18] M. Liberton, R. Howard Berg, J. Heuser, R. Roth, and H. B. Pakrasi,
“Ultrastructure of the membrane systems in the unicellular
cyanobacterium Synechocystis sp. strain PCC 6803,” Protoplasma,
vol. 227, no. 2–4, pp. 129–38, 2006.
[19] N. D. Rawlings, A. J. Barrett, and A. Bateman, “MEROPS: the
database of proteolytic enzymes, their substrates and inhibitors,”
Nucleic Acids Res., vol. 40, no. Database issue, pp. D343–50, 2012.
[20] S. Wickner, “Posttranslational quality control: folding, refolding, and
degrading proteins,” Science, vol. 286, no. 5446, pp. 1888–1893, 1999.
[21] F. U. Hartl, A. Bracher, and M. Hayer-Hartl, “Molecular chaperones in
protein folding and proteostasis,” Nature, vol. 475, no. 7356, pp. 324–
32, 2011.
[22] R. J. Ellis, Molecular Chaperones and Cell Signalling. Cambridge:
Cambridge University Press, 2005.
[23] J. Porankiewicz, J. Wang, and A. K. Clarke, “New insights into the
ATP-dependent Clp protease: Escherichia coli and beyond,” Mol.
Microbiol., vol. 32, no. 3, pp. 449–458, 1999.
[24] S. Langklotz, U. Baumann, and F. Narberhaus, “Structure and
function of the bacterial AAA protease FtsH,” Biochim. Biophys. Acta,
vol. 1823, no. 1, pp. 40–8, 2012.
[25] C. Spiess, a Beil, and M. Ehrmann, “A temperature-dependent switch
from chaperone to protease in a widely conserved heat shock protein,”
Cell, vol. 97, no. 3, pp. 339–47, 1999.
[26] R. Sun, H. Fan, F. Gao, Y. Lin, L. Zhang, W. Gong, and L. Liu, “Crystal
structure of Arabidopsis Deg2 protein reveals an internal PDZ ligand
locking the hexameric resting state,” J. Biol. Chem., vol. 287, no. 44,
pp. 37564–9, 2012.
[27] D. Y. R. Kim, S. C. Ha, N. K. Lokanath, C. J. Lee, H.-Y. Hwang, and K.
K. Kim, “Crystal structure of the protease domain of a heat-shock
protein HtrA from Thermotoga maritima,” J. Biol. Chem., vol. 278,
no. 8, pp. 6543–51, 2003.
30
[28] D. Y. Kim and K. K. Kim, “Structure and function of HtrA family
proteins, the key players in protein quality control,” J. Biochem. Mol.
Biol., vol. 38, no. 3, pp. 266–74, 2005.
[29] J. Huerta-Cepas, S. Capella-Gutiérrez, L. P. Pryszcz, M. Marcet-
Houben, and T. Gabaldón, “PhylomeDB v4: zooming into the plurality
of evolutionary histories of a genome,” Nucleic Acids Res., vol. 42, no.
Database issue, pp. D897–902, 2014.
[30] B. Lipinska, O. Fayet, L. Baird, and C. Georgopoulosl, “Identification,
characterization, and mapping of the Escherichia coli htrA gene,
whose product is essential for bacterial growth only at elevated
temperatures,” J. Bacteriol., vol. 171, no. 3, pp. 1574–1584, 1989.
[31] K. L. Strauch, K. I. T. Johnson, and J. O. N. Beckwith,
“Characterization of degP, a gene required for proteolysis in the cell
envelope and essential for growth of Escherichia coli at high
temperature,” J. Bacteriol., vol. 171, no. 5, pp. 2689–2696, 1989.
[32] T. Clausen, C. Southan, M. Ehrmann, and M. Park, “The HtrA family
of proteases: implications for protein composition and cell fate,” Mol.
Cell, vol. 10, no. 3, pp. 443–455, 2002.
[33] H. Hasselblatt, R. Kurzbauer, C. Wilken, T. Krojer, J. Sawa, J. Kurt, R.
Kirk, S. Hasenbein, M. Ehrmann, and T. Clausen, “Regulation of the
sigmaE stress response by DegS: how the PDZ domain keeps the
protease inactive in the resting state and allows integration of
different OMP-derived stress signals upon folding stress,” Genes Dev.,
vol. 21, no. 20, pp. 2659–70, 2007.
[34] J. Sawa, H. Malet, T. Krojer, F. Canellas, M. Ehrmann, and T. Clausen,
“Molecular adaptation of the DegQ protease to exert protein quality
control in the bacterial cell envelope,” J. Biol. Chem., vol. 286, no. 35,
pp. 30680–90, 2011.
[35] J. Chien, J. Staub, S.-I. Hu, M. R. Erickson-Johnson, F. J. Couch, D. I.
Smith, R. M. Crowl, S. H. Kaufmann, and V. Shridhar, “A candidate
tumor suppressor HtrA1 is downregulated in ovarian cancer,”
Oncogene, vol. 23, no. 8, pp. 1636–44, 2004.
REFERENCES
31
[36] L. Vande Walle, M. Lamkanfi, and P. Vandenabeele, “The
mitochondrial serine protease HtrA2/Omi: an overview,” Cell Death
Differ., vol. 15, no. 3, pp. 453–60, 2008.
[37] S. K. Tanz, I. Castleden, C. M. Hooper, I. Small, and A. H. Millar,
“Using the SUBcellular database for Arabidopsis proteins to localize
the Deg protease family,” Front. Plant Sci., vol. 5, p. 396, 2014.
[38] X. Sun, M. Ouyang, J. Guo, J. Ma, C. Lu, Z. Adam, and L. Zhang, “The
thylakoid protease Deg1 is involved in photosystem-II assembly in
Arabidopsis thaliana,” Plant J., vol. 62, no. 2, pp. 240–9, 2010.
[39] X. Sun, L. Peng, J. Guo, W. Chi, J. Ma, C. Lu, and L. Zhang,
“Formation of Deg5 and Deg8 complexes and their involvement in the
degradation of photodamaged photosystem II reaction center D1
protein in Arabidopsis,” Plant Cell, vol. 19, no. 4, pp. 1347–61, 2007.
[40] P. F. Huesgen, H. Schuhmann, and I. Adamska, “The family of Deg
proteases in cyanobacteria and chloroplasts of higher plants,” Physiol.
Plant., vol. 123, no. 4, pp. 413–420, 2005.
[41] S. Kim, R. A. Grant, and R. T. Sauer, “Covalent linkage of distinct
substrate degrons controls assembly and disassembly of DegP
proteolytic cages,” Cell, vol. 145, no. 1, pp. 67–78, 2012.
[42] L. Truebestein, A. Tennstaedt, T. Mönig, T. Krojer, F. Canellas, M.
Kaiser, T. Clausen, and M. Ehrmann, “Substrate-induced remodeling
of the active site regulates human HTRA1 activity,” Nat. Struct. Mol.
Biol., vol. 18, no. 3, pp. 386–8, 2011.
[43] S. T. Runyon, Y. Zhang, B. A. Appleton, S. L. Sazinsky, P. Wu, B. Pan,
C. Wiesmann, N. J. Skelton, and S. S. Sidhu, “Structural and
functional analysis of the PDZ domains of human HtrA1 and HtrA3,”
Protein Sci., vol. 16, pp. 2454–2471, 2007.
[44] W. Li, S. M. Srinivasula, J. Chai, P. Li, J.-W. Wu, Z. Zhang, E. S.
Alnemri, and Y. Shi, “Structural insights into the pro-apoptotic
function of mitochondrial serine protease HtrA2/Omi,” Nat. Struct.
Biol., vol. 9, no. 6, pp. 436–41, 2002.
[45] J. Kley, B. Schmidt, B. Boyanov, P. C. Stolt-Bergner, R. Kirk, M.
Ehrmann, R. R. Knopf, L. Naveh, Z. Adam, and T. Clausen, “Structural
32
adaptation of the plant protease Deg1 to repair photosystem II during
light exposure,” Nat. Struct. Mol. Biol., vol. 18, no. 6, pp. 728–31,
2011.
[46] W. Sun, F. Gao, H. Fan, X. Shan, R. Sun, L. Liu, and W. Gong, “The
structures of Arabidopsis Deg5 and Deg8 reveal new insights into
HtrA proteases,” Acta Crystallogr. D. Biol. Crystallogr., vol. 69, no. Pt
5, pp. 830–7, 2013.
[47] R. Wrase, H. Scott, R. Hilgenfeld, and G. Hansen, “The Legionella
HtrA homologue DegQ is a self-compartmentizing protease that forms
large 12-meric assemblies,” Proc. Natl. Acad. Sci. U. S. A., vol. 108, no.
26, pp. 10490–5, 2011.
[48] K. Fan, J. Zhang, X. Zhang, and X. Tu, “Solution structure of HtrA
PDZ domain from Streptococcus pneumoniae and its interaction with
YYF-COOH containing peptides,” J. Struct. Biol., vol. 176, no. 1, pp.
16–23, 2011.
[49] M. J. Page and E. Di Cera, “Serine peptidases: classification, structure
and function,” Cell. Mol. Life Sci., vol. 65, no. 7–8, pp. 1220–36, 2008.
[50] H. Czapinska and J. Otlewski, “Structural and energetic determinants
of the S 1 -site specificity in serine proteases,” Eur. J. Biochem., vol.
260, no. 3, pp. 571–595, 1999.
[51] T. Krojer, J. Sawa, E. Schäfer, H. R. Saibil, M. Ehrmann, and T.
Clausen, “Structural basis for the regulated protease and chaperone
function of DegP,” Nature, vol. 453, no. 7197, pp. 885–90, 2008.
[52] T. Krojer, M. Garrido-Franco, R. Huber, M. Ehrmann, and T. Clausen,
“Crystal structure of DegP (HtrA) reveals a new protease-chaperone
machine,” Nature, vol. 416, no. 6879, pp. 455–9, 2002.
[53] T. Krojer, J. Sawa, R. Huber, and T. Clausen, “HtrA proteases have a
conserved activation mechanism that can be triggered by distinct
molecular cues,” Nat. Struct. Mol. Biol., vol. 17, no. 7, pp. 844–52,
2010.
[54] M. Barker, R. de Vries, J. Nield, J. Komenda, and P. J. Nixon, “The
Deg proteases protect Synechocystis sp. PCC 6803 during heat and
light stresses but are not essential for removal of damaged D1 protein
REFERENCES
33
during the photosystem two repair cycle,” J. Biol. Chem., vol. 281, no.
41, pp. 30347–55, 2006.
[55] T. Kieselbach and C. Funk, “The family of Deg/HtrA proteases: from
Escherichia coli to Arabidopsis,” Physiol. Plant., vol. 119, no. 3, pp.
337–346, 2003.
[56] F. Sievers, A. Wilm, D. Dineen, T. J. Gibson, K. Karplus, W. Li, R.
Lopez, H. McWilliam, M. Remmert, J. Söding, J. D. Thompson, and D.
G. Higgins, “Fast, scalable generation of high-quality protein multiple
sequence alignments using Clustal Omega,” Mol. Syst. Biol., vol. 7, p.
539, 2011.
[57] J. Sohn, R. A. Grant, and R. T. Sauer, “OMP peptides activate the DegS
stress-sensor protease by a relief of inhibition mechanism,” Structure,
vol. 17, no. 10, pp. 1411–1421, 2009.
[58] M. Lopo, A. Montagud, E. Navarro, I. Cunha, A. Zille, P. F. de
Córdoba, P. Moradas-Ferreira, P. Tamagnini, and J. F. Urchueguía,
“Experimental and modeling analysis of Synechocystis sp. PCC 6803
growth,” J. Mol. Microbiol. Biotechnol., vol. 22, no. 2, pp. 71–82,
2012.
[59] S. Fulda, F. Huang, F. Nilsson, M. Hagemann, and B. Norling,
“Proteomics of Synechocystis sp. strain PCC 6803,” Eur. J. Biochem.,
vol. 267, no. 19, pp. 5900–5907, 2000.
[60] F. Huang, S. Fulda, M. Hagemann, and B. Norling, “Proteomic
screening of salt-stress-induced changes in plasma membranes of
Synechocystis sp. strain PCC 6803,” Proteomics, vol. 6, no. 3, pp.
910–20, 2006.
[61] F. Huang, E. Hedman, C. Funk, T. Kieselbach, W. P. Schröder, and B.
Norling, “Isolation of outer membrane of Synechocystis sp. PCC 6803
and its proteomic characterization,” Mol. Cell. Proteomics, vol. 3, no.
6, pp. 586–95, 2004.
[62] H. T. Tsui, S. K. Keen, L.-T. Sham, K. J. Wayne, and M. E. Winkler,
“Dynamic distribution of the SecA and SecY translocase subunits and
septal localization of the HtrA surface chaperone/protease during
Streptococcus pneumoniae D39 cell division,” MBio, vol. 2, no. 5, pp.
e00202–11, 2011.
34
[63] A. R. Slabas, I. Suzuki, N. Murata, W. J. Simon, and J. J. Hall,
“Proteomic analysis of the heat shock response in Synechocystis
PCC6803 and a thermally tolerant knockout strain lacking the
histidine kinase 34 gene,” Proteomics, vol. 6, no. 3, pp. 845–64, 2006.
[64] T. Jansén, H. Kidron, H. Taipaleenmäki, T. Salminen, and P.
Mäenpää, “Transcriptional profiles and structural models of the
Synechocystis sp. PCC 6803 Deg proteases,” Photosynth. Res., vol. 84,
no. 1–3, pp. 57–63, 2005.
[65] O. Koksharova, M. Schubert, S. Shestakov, and R. Cerff, “Genetic and
biochemical evidence for distinct key functions of two highly divergent
GAPDH genes in catabolic and anabolic carbon flow of the
cyanobacterium Synechocystis sp. PCC 6803,” Plant Mol. Biol., vol.
36, no. 1, pp. 183–194, 1998.
[66] R. M. Figge, C. Cassier-Chauvat, F. Chauvat, and R. Cerff, “The carbon
metabolism-controlled Synechocystis gap2 gene harbours a conserved
enhancer element and a Gram-positive-like -16 promoter box retained
in some chloroplast genes,” Mol. Microbiol., vol. 36, no. 1, pp. 44–54,
2000.
[67] K. Nakahara, H. Yamamoto, C. Miyake, and A. Yokota, “Purification
and characterization of class-I and class-II fructose-1,6-bisphosphate
aldolases from the cyanobacterium Synechocystis sp. PCC 6803,”
Plant Cell Physiol., vol. 44, no. 3, pp. 326–33, 2003.
[68] T. Ogawa, E. Marco, and M. I. Orus, “A gene (ccmA) required for
carboxysome formation in the cyanobacterium Synechocystis sp.
strain PCC6803,” J. Bacteriol., vol. 176, no. 8, pp. 2374–8, 1994.
[69] C. Trautner and W. F. J. Vermaas, “The sll1951 gene encodes the
surface layer protein of Synechocystis sp. strain PCC 6803,” J.
Bacteriol., vol. 195, no. 23, pp. 5370–80, 2013.
[70] A. El Zoeiby, F. Sanschagrin, and R. C. Levesque, “Structure and
function of the Mur enzymes: development of novel inhibitors,” Mol.
Microbiol., vol. 47, no. 1, pp. 1–12, 2002.
[71] C. R. Sweet, A. H. Williams, M. J. Karbarz, C. Werts, S. R. Kalb, R. J.
Cotter, and C. R. H. Raetz, “Enzymatic synthesis of lipid A molecules
with four amide-linked acyl chains. LpxA acyltransferases selective for
REFERENCES
35
an analog of UDP-N-acetylglucosamine in which an amine replaces
the 3"-hydroxyl group,” J. Biol. Chem., vol. 279, no. 24, pp. 25411–9,
2004.
[72] E. Sauvage, F. Kerff, M. Terrak, J. a Ayala, and P. Charlier, “The
penicillin-binding proteins: structure and role in peptidoglycan
biosynthesis,” FEMS Microbiol. Rev., vol. 32, no. 2, pp. 234–58, 2008.
[73] M. Marbouty, K. Mazouni, C. Saguez, C. Cassier-Chauvat, and F.
Chauvat, “Characterization of the Synechocystis strain PCC 6803
penicillin-binding proteins and cytokinetic proteins FtsQ and FtsW
and their network of interactions with ZipN,” J. Bacteriol., vol. 191,
no. 16, pp. 5123–33, 2009.
[74] S. Yoshihara, X. Geng, S. Okamoto, K. Yura, T. Murata, M. Go, M.
Ohmori, and M. Ikeuchi, “Mutational analysis of genes involved in
pilus structure, motility and transformation competency in the
unicellular motile cyanobacterium Synechocystis sp. PCC 6803,”
Plant Cell Physiol., vol. 42, no. 1, pp. 63–73, 2001.
[75] D. Bhaya, A. Takahashi, P. Shahi, and R. G. Arthur, “Novel motility
mutants of Synechocystis Strain PCC 6803 generated by in vitro
transposon mutagenesis,” J. Bacteriol., vol. 183, no. 20, pp. 6140–
6143, 2001.
[76] H. Miranda, O. Cheregi, S. Netotea, T. R. Hvidsten, T. Moritz, and C.
Funk, “Co-expression analysis, proteomic and metabolomic study on
the impact of a Deg/HtrA protease triple mutant in Synechocystis sp.
PCC 6803 exposed to temperature and high light stress,” J.
Proteomics, vol. 78, pp. 294–311, 2013.