Download protein phosphorylation in bacteria

PROTEIN PHOSPHORYLATION IN BACTERIA
Michał Obuchowski, PhD
Laboratory of Molecular Bacteriology
Department of Medical Biotechnology of IFB UG-MUG
Course book prepared as part of the project: „Kształcimy najlepszych kompleksowy program rozwoju
doktorantów, młodych doktorów oraz akademickiej kadry dydaktycznej Uniwersytetu Gdańskiego”
Project no: UDA-POKL.04.01.01-00-017/10-00
Intercollegiate Faculty of Biotechnology UG-MUG
Gdańsk 2011
Table of contents:
Foreword .................................................................................................................................... 5
Introduction – protein phosphorylation is everything ............................................................... 6
1. Chemotaxis, a very important question: where to swim? ................................................... 13
2. General stress response – How to manage stress?.............................................................. 15
3. Control of nitrogen assimilation ........................................................................................... 17
4. Many stimuli for one response of Vibrio cholerae ............................................................... 19
5. A very important question – to sporulate or not to sporulate?........................................... 21
6. Bioluminescence – in a mass we will shine like stars ........................................................... 29
7. Genetic competence – to be competent or not? ................................................................. 32
8. PTS – a matter of taste ......................................................................................................... 35
Literature .................................................................................................................................. 39
Protein phosphorylation in bacteria
by Michał Obuchowski
3
Foreword
The origin of studying living organisms dates back to the moment, when humans started to
develop a culture, over 19 000 years ago. The first record of it is domestication of dogs. People
accumulated enough information to consider that these animals might be useful. Later, the amount
of possessed knowledge increased along with capabilities of humans. Now, we are able to observe
molecular details of different aspects of life, but we face another problem – an enormous amount
of information necessary to describe even a single cell organism at detailed molecular level. Below
you will find some selected information about basis of control by phosphorylation and several
examples of it utilized by bacteria. The phenomenon of reversible phosphorylation of Ser/Thr/Tyr
was discovered in 1956 and for quite long scientist were convinced that it is specific only for
eukaryotes. Only in the last several years it became clear that protein phosphorylation also plays
a very important role in bacteria. I would also like to comment on the common belief that bacteria
are simple and primitive organisms. It is doubtless that bacteria are the simplest unicellular
organisms on Earth. Nevertheless, complexity of functioning and regulation of these organisms still
extends far beyond our knowledge. In the following sections you will have a chance to see the true
character of these “primitive and simple” organisms. I hope that the selected examples of different
regulatory networks described later will give you at least a glimpse of the general view of regulatory
cycles and intersections between different aspects of cell physiology and behaviour.
Protein phosphorylation in bacteria. Foreword
by Michał Obuchowski
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Introduction – protein phosphorylation is everything
Living organisms need to conduct several hundred of different chemical reactions at the
same time. In this process the necessary tools are the enzymes which catalyse particular reactions.
In order to manage such abundance of different functional proteins, living cells had to develop a set
of mechanisms allowing for control of their activity. This task is achieved at various levels
of expression of genetic information, from chromatin structure to direct control of enzyme activity in
the cell. This last step, direct control of enzymatic activity is the most important one, because it is
subject to instantaneous changes due to different intra and/or extracellular stimuli. Even at this level
of control living cells develop different ways of controlling enzymatic activity. One of the most
commonly used is modification by phosphorylation. This way of control of enzymatic activity has
several advantages in comparison with other methods:
i) the phosphate group is easy accessible in the cell;
ii) the requirements for phosphorylation site are permissive, i.e. any protein theoretically
possesses several sites of phosphorylation;
iii) the phosphate group exhibits high charge density at physiological pH;
iv) the addition of such a group to a peptide or a protein can be easily reversed;
v) the control cycle based on this mechanism requires only two enzymes: one for introducing the
phosphate group into the proteins (a protein kinase), another for removing it (a protein
phosphatase) (Johnson, et al., 2001).
When considering accessibility of the phosphate group, which can be used for protein phosphorylation, adenosine triphosphate (ATP) is an obvious candidate. This molecule is present in cells
of all living organisms. Moreover, it is the molecule most commonly used as energy donor for
biocatalysis and the release of energy connected with phosphate bond. Obviously, ATP is not the
only phosphate donor for protein kinases. Several kinases which can use GTP instead of ATP have
been described. In addition, few other small molecules, such as acetyl phosphate, can be used
as phosphate donors.
If we compare elements necessary for the binding site of the regulatory molecule
(e.g. lactose for LacR protein) and the phosphorylation site, this second turns out to be very simple
because the presence of hydroxyl, amine, carboxyl or sulfhydryl groups are sufficient conducting of
this reaction. In a standard set of twenty amino acids nine can be theoretically phosphorylated
(Duclos, et al., 1991). Five of these are commonly phosphorylated.
It is really hard to imagine a protein molecule which does not posses any serine, threonine,
tyrosine, histidine or aspartic acid residue. Based on that we can assume that any protein can be
a potential substrate for a protein kinase. Of course, the presence of phosphorylable residue alone is
not a sufficient factor without proper surroundings, but this is still very simple, if we compare it to for
instance the structure of binding cleft for regulatory molecules. If phosphorylation may lead to
change of enzymatic activity, it must be able to force some changes in the target molecule. It can be
achieved in at least two ways: blocking of the catalytic cleft of target enzyme or changing the
structure of this enzyme. In fact, both methods are used. An example of such mechanisms
is characteristic for citrate synthase, where the addition of phosphate group blocks accessibility of
catalytic cleft for the substrate. Nevertheless, the second way is more commonly used. Introduction
of a phosphate group to the enzyme molecule may change its structure in two ways: a physical
change of its shape and addition of electrostatic charge. Usually both effects act synergistically.
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1TProtein phosphorylation in bacteria. Introduction – protein phosphorylation is everything
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Efficient regulation of enzymatic activity also needs to be reversible. Removal of the phosphate
group from protein molecule is achieved by catalysed hydrolysis resulting in regeneration of acceptor
site and a molecule of phosphoric acid. It is worth of pointing out that a cell requires only two
enzymes to maintain regulation by reversible protein phosphorylation: a protein kinase, which adds
a phosphate group to protein molecules, and a protein phosphatase, which removes it. If we realize
that such kinase-phosphatase couples can exhibit activity towards many different enzyme molecules
in the cell, the total cost of this type of control turns out to be relatively low. Taking all this into
account, it is not surprising that the control of protein function by reversible phosphorylation is the
most commonly used way of activity regulation found in all living organisms.
As it was said phosphorylation status of a protein influences not only its activity, but also its
interaction with other proteins in the cell, as well as its localisation. Analysis of human proteome
revealed that almost 1/3 of proteins are phosphorylated. In order to manage such a vast amount of
proteins, human genome encodes 530 probable protein kinases and 130 protein phosphatases
(Hunter, 1995) (Hrabak, et al., 2003). In prokaryotic organisms many processes are also regulated by
protein phosphorylation. Although members of the eukaryote kingdom have several hundred of
protein kinases, some of bacteria have a dozen or so of such enzymes. Similarly to higher organisms
reversible phosphorylation of proteins plays a crucial role in many cellular processes, such as basic
metabolism regulation, horizontal gene transfer, virulence, sporulation, etc. Bacterial signal
transduction pathways are usually much simpler than eukaryotic ones.
Firstly, we must consider which amino acids may be phosphorylated. The obvious triad is
serine, threonine and tyrosine. These amino acids possess side chains containing a hydroxyl group,
which can be phosphorylated. Its phosphor-derivatives are O-phosphates or O-phosphomonoesters.
Properties of phosphoserine (pSer or pS) and phosphothreonine (pThr or pT) are very similar. These
compounds are stable at physiological pH, resistant to acid, hydroxylamine and pyridine treatment.
Even more stable is phosphotyrosine (pTyr or pY). This phosphoric derivative is also resistant to
alkali, in addition to previously described features. Theoretically an additional amino acid is available,
which possesses a hydroxyl group – hydroxylproline. However, no phosphohydroxylproline has been
found in proteins so far. O-phosphates are characteristic for “eukaryotic-type” of protein
phosphorylation. But, if you thought that O-phosphate are absent in prokaryotic cells, you would be
wrong. About twenty years ago O-phosphomonoesters were discovered in bacterial cells and
currently no one negates their presence. The next group of phosphor-derivatives which has been
found inside living cells are N-phosphates or phosphoramidates. This group is produced by
phosphorylation of the basic amino acids: arginine, histidine and lysine. N-phosphates are much
more labile at physiological pH than O-phosphates. They are resistant only to alkali treatment (with
exception of phosphoarginine). The third group of phosphoamino acids are acylphosphates,
or phosphate anhydrides. These derivatives are generated by phosphorylation of acidic amino acids:
aspartic acid and glutamic acid. Acylphosphates are unstable at physiological conditions. The last,
fourth group of phosphoamino acids, which is relatively less abundant, are S-phosphates,
or thioesters. This group contains only one amino acid which can be phosphorylated – cysteine.
Phosphocysteine is the most stable phosphor-derivative of all listed above. It is resistant to acid,
alkali, hydroxylamine and pyridine treatment. Two types of phosphoproteins are known to be
present in the living cells: enzymes that are transiently phosphorylated, usually at their active sites,
and proteins that are phosphorylated by protein kinases (Duclos, et al., 1991). The first group of
phosphoproteins is not in our area of interest and we will focus on the second one below.
Protein phosphorylation in bacteria. Introduction – protein phosphorylation is everything
by Michał Obuchowski
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Phosphoproteins are products of enzymes, which transfer orthophosphate group from its
donor to acceptor site located in the side chain of the amino acid residue. Generally the donor is γphosphate of ATP or another nucleoside triphosphate, but individual enzymes may have other
phosphate donors. Because most of the kinases have multiple substrates its classification is based on
the specificity of acceptor amino acid. In accordance with it, the Nomenclature Committee of the
International Union of Biochemists has recommended the following names:
• Phosphotransferases with a hydroxyl group as an acceptor (i.e. serine or threonine), called
protein serine/threonine kinases (E.C. 2.7.10), which produce O-phosphates;
• Phosphotransferases with a phenolic group as an acceptor (i.e. tyrosine), called protein tyrosine
kinases (E.C. 2.7.11), which produce also O-phosphates;
• Phosphotransferases with amine or imine group as an acceptor (i.e. arginine, lysine or histidine),
called protein histidine kinases (E.C. 2.7.12), which produce N-phosphates;
• Phosphotransferases with sulfhydryl group as an acceptor (i.e. cysteine), called protein cysteine
kinases (E.C. 2.7.13), which produce S-phosphates;
• Phosphotransferases witch acyl group as an acceptor (i.e. aspartic acid or glutamic acid), called
protein aspartyl or glutamyl kinases (E.C. 2.7.14), which produce acyl-phosphates.
Enzymes of the first two classes are well known, whereas the other classes have been less
characterized. Advances in the DNA sequencing techniques lead to fast increase in the number of
known genomes resulting in a very rapid increase in the number of putative protein kinases
described in different organisms. When phosphoproteins are generated inside a cell, there must be
a possibility of reversing this process, i.e. dephosphorylate the phosphoprotein. In order to do this,
the cell possesses a set of specialized enzymes, called protein phosphatases. This functional group of
enzymes is diverse in terms of the evolutionary ancestor, its mechanism of catalysis, and the number
of described members. In general, protein phosphatases are divided on the basis of its substrate
specificity into serine/threonine and tyrosine phosphatases. The first group is subdivided into PPP
and PPM family. The second consists of classical PTP phosphatases, low molecular weight tyrosine
phosphatases (LMW PTP) and dual specificity PTP (Shi, et al., 1998). The PPP family of
serine/threonine phosphatases is the most quantitatively significant source of protein phosphatase
activity in eukaryotic cells. It includes PP1, PP2A and PP2B related phosphatases. Usually cells of
higher organisms contain a limited number of catalytic subunits, which are controlled by association
with a large family of regulatory and targeting subunits, which temporally and spatially control its
action (Cohen, 1991). During analysis of known prokaryote genomes several ORFs encoding putative
PPP phosphatase have been described, few of which have been described at biochemical and
functional level. There are typical examples of serine/threonine phosphatases in this set of described
enzymes, as well as a few other, which can dephosphorylate all O-phosphates. The next family, PPM,
includes serine/threonine protein phosphatases which are characterized by absolute requirement for
divalent metal ion necessary for its catalytic function. In contrast to PPP family, which members are
found and described in all high-rank taxa, the PPM are absent in archebacteria. It includes eukaryotic
PP2C and the puruvate dehydrogenase phosphatase, along with many bacterial ones. Inside this
family, PPM members are postulated to be a part of different signalling pathways, which respond to
environmental stress (Kennelly, et al., 1999). The last group includes protein phosphatases, which
dephosphorylate tyrosine residues. In contrast to many PPP and all PPM phosphatases, all tyrosine
phosphatases are independent of the presence of metal ions. Moreover, a covalently bound
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1TProtein phosphorylation in bacteria. Introduction – protein phosphorylation is everything
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intermediate enzyme-substrate is created during dephosphorylation reaction. These enzymes are
divided into three families: tyrosine phosphatase (PTP), low molecular weight tyrosine phosphatases
(LMW PTP), and dual specific PTP (dsPTP). The first family includes classical tyrosine phosphatases.
In the catalytic cleft of these enzymes there is a conserved cysteine residue, which is crucial for
enzymatic activity. Those enzymes have molecular mass of about 30-40 kDa. Second group, Low
Molecular Weight tyrosine phosphatases (LMW PTP), consists of small molecules of molecular mass
equal to more or less 15 kDa. Both families described above are specific for phosphotyrosine only.
The last group includes enzymes, which are able to use all O-phosphates as substrates. Dual
specificity should be interpreted as ability to dephosphorylate phosphoserine/phosphotreonine,
as well as phosphotyrosine (Shi, et al., 1998). In general, protein phosphatases are more diverse than
protein kinases (Kennelly, 2001) (Bhaduri, et al., 2005). As mentioned above, eukaryotic cells may
possess over hundred enzymes of this class. Prokaryotic organisms usually posses few of them, but
a system based on phosphorylation of histidine and aspartic acid residues, generally called two
component system, plays an important role in these organisms (Saito, 2001). As the name suggests,
these signalling pathways may consists of only two proteins: a sensor histidine kinase and a response
regulator. Sensor kinase is usually a membrane linked protein with extracellular or periplasmatic
sensory domain, which receives the stimuli. Next, the signal is transferred through the kinase
molecule and affects activity of the kinase domain (Parkinson, 1995). There is no general rule
regarding the effect of the stimuli: some kinase domains become active, some become inactive after
receiving the signal. Let’s assume that reception of the signal increases the activity of the kinase
domain. In response to the stimulus autophosphorylation occurs on conserved histidine residue of
the kinase. It is worth of noticing that autophosphorylation of the kinase histidine is fully reversible.
Next, the phosphate group is transferred to aspartic acid residue of the response regulator molecule.
Then kinase returns to unphospohrylated state and is ready for the next cycle of the reaction.
Phosphorylated response regulator diffuses into the cell and binds its molecular target, e.g. a DNA
sequence or partner protein (Stock, et al., 1995). At this stage the two component system is
considered to have been turned on. In order to maintain cell homeostasis, it is crucial to shut off the
action of response regulator when the stimulus is no longer present. The system can be turned off in
a few different ways. Firstly, receptor kinase without the ligand may act as a phosphatase, which
results in removal of phosphate from the response regulator molecule. Secondly, response regulator
may spontaneously undergo autodephosphorylation. Thirdly, some two component systems are
composed of specialized protein phosphatases, which dephosphorylate response regulator
molecules. Fourthly, it has already been shown that some proteins, which bind to the phosphorylated response regulator, stimulate its autophosphatase activity, but this additional protein is not
a protein phosphatase. Fifthly, proteins similar to the response regulator may exist in the cell and
they can remove phosphate from it, becoming phosphorylated; however, these molecules cannot act
as response regulators. All described possibilities may act in parallel in different two component
signalling pathways. Depending on cellular needs, cells use different architecture of those systems,
which may consist of several histidine kinases or phosphatases. However, even such systems are
called two component systems.
Let’s take a closer look at conserved motifs found in protein kinases and phosphatases. Let us
start with kinases. Histidine kinase has a conserved core, which extends to over 200 residues. This
part of histidine kinase has ATP binding motif and is able to catalyze phosphorylation of a side chain
of histidine residue. The important catalytic part is always surrounded by non-conserved regions,
Protein phosphorylation in bacteria. Introduction – protein phosphorylation is everything
by Michał Obuchowski
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which perform specific regulatory functions. Histidine residue, which undergoes phosphorylation,
is generally located inside the catalytic part of the kinase. The catalytic domain is subdivided into two
parts: N-subdomain, containing conserved histidine residue, and C-subdomain, containing four highly
conserved regions called N, D, F and G boxes. These motifs are involved in tertiary structure of the
protein, which is responsible for binding of nucleotides. Histidine and serine/threonine kinases have
two subdomains, functions of which are inverted. N-subdomain of histidine kinases (C-subdomain
in serine/threonine kinases) is responsible for substrate binding, whereas C-subdomain binds the
Mg-ATP (N-subdomain in serine/threonine kinases) complex. Phosphorylation of conserved histidine
does not proceed by an intramolecular mechanism. It has been demonstrated that one kinase
monomer catalyzes phosphorylation of a histidine residue in the second monomer. It means that
functional complex of histidine kinase must be a dimer. In most histidine protein kinases the
phosphorylated histidine is located around 100 amino acids from N box towards the N-terminus,
close to one of the ends of conserved catalytic domain. Mutations in this region usually result in
complete loss of kinase activity. However, a histidine kinase lacking conserved histidine residues has
been described. This enzyme is well characterized and phosphorylates the serine residue of
a substrate molecule. Histidine kinases have been described in bacteria, as well as in yeast and
plants. Studies of the regulation of autophosphorylation rate revealed that mechanism is based on
control of transition of monomer to dimer (Stock, et al., 1995).
Now let us discuss serine/threonine kinases. In this group of enzymes the catalytic domain
consists of about 260 residues. Similarly to histidine kinases, the catalytic domain is divided into two
lobes: small and large. The small lobe consists of several antiparallel β-sheets and two helical regions.
Its function is binding the Mg-ATP complex. The large lobe consists mainly of α-helices, with a small
fraction of β-sheets. This part of kinase molecule is associated with catalysis and peptide binding.
The most important region of the serine/threonine kinase is located between the lobes, and it is
usually called the catalytic cleft (Taylor, et al., 1996). Comparative studies of amino acid sequences of
numerous serine/threonine protein kinases revealed the existence of several conserved motifs,
characteristic for these enzymes. Motifs I–III are located in the small lobe, while the fourth motif is
a linker between the lobes. Motifs V–XI are located in the large lobe of the kinase. Glycine-rich loop,
homologous to the G-box of histidine kinase is part of motif I. Next very important residue, lysine
which interacts with the phosphate chain of ATP, is placed in motif II. Mutation of conserved lysine
residue usually abolishes enzymatic activity of the kinase. Another important residue involved in
nucleotide binding is glutamic acid, which is placed in motif III. Its side chain together with
mentioned lysine interacts with phosphate chain of ATP, as well as Mg2+ ions, in order to ensure
proper localisation and orientation of γ-phosphate for phosphotransfer reaction. The role of the large
lobe in substrate binding is less clear, because it needs to be partially shared by all substrates.
Approaches for establishing the consensus sequence for phosphorylation site have been unsuccessful
and all attempts to predict such a site with high probability are questionable. It is known that the
substrate occupies the cleft between the two lobes and interacts with several residues of the kinase
molecule. The large lobe contains conserved motifs from VIa–XI. The DxKxxN motif placed in motif
VIb, together with motifs VIII, is involved in proper orientation of the substrate. In addition, lysine
residue from motif VIb plays an important role during phosphotransfer event. Some authors named
motif VIb as the catalytic loop. Conserved triplet DFG from motif VII is involved in orientation of
γ-phosphate. Motif IX stabilizes position of the catalytic loop. Precise role of motifs X and XI
is unknown. In addition to conserved motifs, there is a region in the serine/threonine kinase
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1TProtein phosphorylation in bacteria. Introduction – protein phosphorylation is everything
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molecule, which deserves special attention: activation loop. It is placed between motif VII and VIII,
has no defined secondary structure nor any conserved residues. However, in many kinases this motif
plays a crucial role in the regulation of kinase activity. The catalytic loop often contains one or more
residues undergoing autophosphorylation, which has great influence on the activity status of the
particular kinase (Shi, et al., 1998) (Vulpetti, et al., 2004). The catalytic subunits of serine/threonine
kinase are usually a part of the bigger molecule, with one or more domains regulating kinase activity
of the whole molecule. It should be pointed out that many of serine/threonine kinases undergo
autophosphorylation, but of a very different nature from the histidine kinase. In case of histidine
kinase, autophosphorylation is absolutely crucial for transfer of phosphate to the substrate.
Serine/threonine kinases undergo autophosphorylation solely for regulatory purposes. Transfer of
γ-phosphate from the donor to the target is always direct.
The second side of the coin are protein phosphatases. Many regulatory circuits based on
histidine kinase lack a dedicated phosphatase. However, some of them include auxiliary aspartate
phosphatases. Up to date, four different families of auxiliary phosphatases have been described –
CheZ, CheC/CheX/FliY, Spo0E, and Rap. Unfortunately, molecular details of the catalytic mechanisms
of only the first two families are known (Silversmith, 2010). Generally speaking, hydrolysis of the
phosphate group from aspartyl residues involves an attack of a nucleophilic water molecule on the
phosphorous atom, where aspartic acid carboxylate is the leaving group. Data collected so far
suggests that despite different topologies, aspartyl phosphatases from various families may utilize
a similar mechanism in order to catalyze dephosphorylation of the response regulator. In contrast to
two component systems, all pathways based on O-phosphates require the presence of specific
protein phosphatases. In addition to the characteristics of O-phosphatases described above, PPP
family has three conserved motifs, which are separated by several amino acids (GDXHG, GDXXDRG,
GNHE). Usually, catalytic subunits of PPP phosphatase span a region of approximately 220 amino
acids. The conserved motifs are shared by PPP phosphatases with diadenine tetraphosphate
hydrolases, which may suggest the existence of a common ancestor. The next family, PPM
phosphatases, have a catalytic domain, which is about 290 amino acids long. Twelve conserved
regions (1–4, 5a, 5b, and 6–11) are present inside this domain, with only 8 well conserved amino
acids. The most important four aspartic acid residues are placed in regions 1, 2, 8 and 11. These
coordinate divalent metal ions required for enzymatic activity. The last group of O-phosphatases,
tyrosine phosphatases, share characteristic active site signature motifs, CXXXXXR. A conventional PTP
catalytic subunit is approximately 250 amino acid residue long, in contrast to LMW PTPs, which
consist of 140 amino acids or less. CXXXXR motifs is placed in the centre of conventional PTPs,
or close to the N-terminus in LMW PTPs. All tyrosine phosphatases employ catalytic mechanisms
(Shi, et al., 1998).
After this brief characterisation of protein kinases and phosphatases let us take a closer look
at substrates. Response regulators (RR), which are phosphorylated by histidine kinases, share
a structure similar to that of the recipient domain, which possesses a conserved aspartic acid residue.
The size of this domain is usually about 120 amino acids and this domain is connected via a linker
sequence to effector domain, if this exists. In some cases, the recipient domain of RR protein is linked
to the histidine kinase at C-terminus. The most extensively studied response regulator is CheY
protein. This molecule is the recipient domain only and its function is to switch the direction of
rotation of the flagellar motor during chemotactic movement of bacteria. This protein was
crystallized for the first time in late 80s (Stock, et al., 1993). It consists of the central bundle of
Protein phosphorylation in bacteria. Introduction – protein phosphorylation is everything
by Michał Obuchowski
11
β-sheets, which is surrounded by α-helices. Sequences of other response regulators confirm this
structure. The phosphorylation site is placed at the aspartic acid residue 57, located between β3 and
α3. Localization of the phosphorylated residue in CheY is conserved in other RRs. In addition to the
recipient domain, most of RR proteins possess an effector domain. It has been demonstrated that the
recipient domain in unphosphorylated state exhibits an inhibitory effect on the effector domain and
that phosphorylation counteracts this inhibitory effect. In the light of this information, CheY is an
unusual response regulator, because it does not contain any effector domain. The phosphorylated
form of CheY binds to the flagellar switch apparatus, FliM protein. Phosphorylation of CheY causes
global restructuring, which results in significant perturbations on the opposite end of the molecule.
However, precise description of differences between active and inactive forms of CheY still remains
unclear. CheY and its function in chemotactic signal transduction is described in more detail below.
Unexpectedly, we can not say anything precise about proteins, which are substrates for
O-phosphorylation, except that they have serine, threonine or tyrosine residues. Possessing at least
one of these amino acids is the only connection. As mentioned above, nearly every protein with
hydroxyl group in the side chain can be phosphorylated or dephosphorylated. Their structures are
too diversified to lead to any conclusions. Considering this, currently no tools are available, which
would allow to predict, if a particular molecule may be a substrate for protein kinase or phosphatase.
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1TProtein phosphorylation in bacteria. Introduction – protein phosphorylation is everything
by Michał Obuchowski
1. Chemotaxis, a very important question: where to swim?
Some bacteria are capable of active movement in their environment. This movement can be
realized by several methods: swimming, swarming, gliding, tumbling, or twitching. The fastest and
most spectacular method is obviously swimming. This type of movement allows bacteria to reach the
speed of over 20 micrometers per second (Chattopadhyay, et al., 2006) (Amsler, et al., 1993), which
is a very high speed. If we converted it to human frame of reference, it would be about 70km/h. Why
do bacteria need to move? The answer is simple: in order to search for a better environment. When
bacteria are inoculated to the fresh medium, they are immotile. The fresh medium is rich in nutrients
and there is no need to move. When a culture continues to grow, conditions change for worse. Close
to entry into the stationary phase bacteria become motile and try to find a better place to grow.
The question is: how can bacteria monitor the environment and use such information for choosing
the direction of swimming? In order to achieve this, bacteria possess a specialized group of
receptors, called chemotactic receptors. These sensors are localized together at one of the cell poles
and their coordinated actions serve as a bacterial “nose”. Information received by these receptors
includes concentration of oxygen, sugars, amino acids, peptides and probably several other small
molecules (Szurmant, et al., 2004). Stimuli received by receptors are transmitted to the two
component system, CheA-CheY. Then, the signal is passed to flagellar motor and the bacterium can
swim. The whole system looks very simple. However, we have to consider the nature of taxis.
By definition, it is a directional movement. This type of movement can be observed in many different
organisms: protista, insects, animals or plants, though bacterial chemotaxis seems to be a bit of overinterpretation. Bacteria have two styles of movement: smooth swimming, when a cell moves straight
forward, and tumbling, when a cell rotates randomly at the same place. Signals received from
chemoreceptors result only in a change of time periods of smooth swimming and tumbling.
In general, this is sufficient for a bacterial cell to advance in chosen direction, but particular events of
smooth swimming occur in randomly selected directions. How can bacteria employ this, a bit chaotic,
movement? The key element is transfer of stimuli from receptors to histidine kinase, CheA. When
bacteria is moving in the expected direction (towards a better environment), receptors sense
an increase of stimuli, e.g. increased concentration of glucose. This leads to transmission of
appropriate signals to CheA kinase and a decrease in its activity. Lowering the concentration of
phosphorylated response regulator, CheY, delays switching of movement from smooth movement to
tumbling, and the bacterium continues moving forward. At the same time receptors undergo
modification by methylation, which changes its sensitivity to stimuli. A very simple mechanism of
memory of the bacterial cell in changing receptor sensitivity by its modification. After a tumbling
event, which must occur even when bacteria move in the expected direction, the cell starts smooth
swimming again. If the direction is still proper, CheA kinase is kept at a low level of activity. In the
opposite situation receptors sense a decrease of stimuli. This change is transmitted to CheA kinase
and its activity rises. If more CheY is phosphorylated in the cell, smooth swimming step is shortened
and replaced by a tumbling event. By regulating the frequency of tumbling in this way, bacteria can
approach an attractant or avoid a repellent. This looks quite simple, does it not? Yes, but it is not so
simple, when we look into the details of molecular interactions between receptors and all proteins
involved in transmission of signals to the flagellar motor. cheA gene, coding for histidine kinase,
produces two proteins: CheAL, which is a full-length kinase, and CheAS, which is a short form, devoid
of conserved histidine residue (Matsumura, et al., 1977). Both versions of CheA kinase form the
Protein phosphorylation in bacteria. 1. Chemotaxis, a very important question: where to swim?
by Michał Obuchowski
13
CheAL- CheAS heterodimer, in which trans-phosphorylation occurs (Swanson, et al., 1993). Proper
transmission of chemotactic signals from receptors to the kinase complex requires another protein,
CheW. This protein forms a stable complex with CheAL and CheAS, in equimolar ratio (Matsubara, et
al., 1990). CheW protein is also necessary for chemotactic receptor aggregation and association of
the kinase dimer with receptors. CheA alone does not interact with receptors. Processing the signal
requires passing of the phosphate group to the response regulator, CheY. CheY can interact with
CheAL alone, with a CheAL- CheAS complex, or with a CheAL- CheAS-CheW complex, towards which
it exhibits highest affinity (McNally, et al., 1991). After phosphorylation of CheY (p-CheY), it loses the
affinity to CheA-CheW complex and can diffuse to flagellar switch. The molecular target of p-CheY
is FliM or FliG – interaction with one of these proteins causes reversal of rotation from CCW (counter
clockwise) to CW (clockwise) and results in tumbling (Welch, et al., 1993). In other words, p-CheY is a
tumbling agent. This type of movement should last for a very short period of time, so a possibility of
removing p-CheY protein is required. In order to achieve this, cells use CheZ protein, which is not a
protein phosphatase. p-CheY undergoes autodephosphorylation, the rate of which is accelerated by
its interaction with CheZ (Hess, et al., 1988). Moreover, CheZ can form a complex with CheAS, but not
with CheAL. Such a complex stimulates dephosphorylation of p-CheY faster than CheZ alone. The next
part of chemotactic signal processing is connected with methyltransferase CheR and methylesterase
CheB, proteins which respectively add and remove methyl group from the chemotactic receptor
(Stewart, et al., 1987). CheR protein is constitutively active. The rate of metylation of receptors is
regulated by the presence of stimuli and it results in a conformational change. In case of CheB the
situation is different. CheB undergoes phosphorylation by CheA kinase and it is more active in that
state. Motility and chemotaxis are a part of global stress response network. Because of this, several
other factors influence expression of Che and flagellar genes, e.g. increased intracellular
concentration of cAMP stimulates chemotactic behaviour (Komeda, 1982). Although chemotaxis is
one of the best understood signal transduction pathways, several details of this mechanism still
remain unknown, like molecular details of transmission of signals from a receptor to CheA kinase, the
way p-CheY causes reversal of flagellar rotation, or the way CheZ influences the interaction between
p-CheY and switch proteins. In addition, several functional differences between E. coli chemotactic
machinery and other species of bacteria have been described, e.g. the mode of swimming is also
coupled with sugar phospho-transport (PTS). In E. coli a signal coming from PTS system is transferred
directly to CheA kinase and lowers its activity. As a result, intracellular concentration of p-CheY drops
and cell swims smoothly for a longer time. This mechanism is called “receptor independent”
(Szurmant, et al., 2004). Different situation can be found in B. subtilis. In this bacterium, a signal
coming from PTS systems is passed through chemotactic receptors. In response to this stimulus
receptors stimulate the kinase activity of CheA, which leads to an increase of p-CheY in the cell
and smooth swimming again. How is it possible? p-CheY protein in E. coli and B. subtilis exhibit
opposite effects on flagellar switch. In the first bacterium, p-CheY stimulates changing of direction of
rotation from CCW to CW. In the second bacterium, elevated concentration of phosphorylated
response regulator decreases the frequency of CCW to CW rotation switch, causing smooth
swimming (Ordal, et al., 1993).
14
1TProtein phosphorylation in bacteria. 1. Chemotaxis, a very important question: where to swim?
by Michał Obuchowski
2. General stress response – How to manage stress?
Bacteria can be found in any terrestrial environments, from the poles to volcano funnels
found deep in the ocean. To survive in such varied conditions, microorganisms have developed
a wide spectrum of permanent or temporary adaptations. If the first profile can be considered to be
properties of particular species, then the second profile, which can be present only in certain
conditions, should be called stress response. An example of it can be soil bacterium Bacillus subtilis.
From the perspective of this very small organism, the soil environment is full of sharp gradients:
temperature, humidity, availability of oxygen and nutrients. Moreover, the surroundings are full of
different micro and macro organisms. It is postulated that 1 gram contains over 108 living bacteria
and it is well known that other species sharing the same environment are often competitors. To be
up to this task, bacteria possess various kinds of stress responses, which allow for temporal
modifications of biochemical properties, like when bacteria are exposed to non-optimal temperature,
the cell synthesizes a set of highly specialized proteins (heat or cold shock proteins), which allow it to
survive. B. subtilis has a highly specialized set of proteins called rsb (regulators of sigma B), which are
used for sensing and switching stress response. The key element of rsb regulon is sigB gene, which
encodes an alternative sigma subunit of the RNA polymerase (Handelwag, et al., 1979). Additional
components of general stress response have been indentified later. Currently, it is known that
at least 17 rsb genes are involved in control of activity of sigmaB and that a general stress response
regulon consists of 150 other genes (Hecker, et al., 2007). The core of the system consists of SigB and
two other proteins: RsbW and RsbV. RsbW is also known as an anti-sigmaB factor. This protein forms
a stable complex with SigB in vivo, in which SigB is inactive. In addition, RsbW is a protein kinase,
of which the only known substrate is RsbV. This protein in named an anti-anti-sigmaB factor, because
it can remove SigB from SigB-RsbW complex and bind RsbW. When RsbV is bound to RsbW kinase,
SigB protein becomes active and can form a complex with the core of RNA polymerase. Regulation of
RsbV activity is based on phosphorylation by RsbW and dephosphorylation by RsbU or RsbP protein
phosphatases. When RsbV is dephosphorylated, SigB is released from the complex, which results in
activation of general stress response, while the phosphorylated form is inactive (Price, 1993). Protein
phosphatases RsbU and RsbP are responsible for dephosphorylation of RsbV in response to
environmental or energetic stress, respectively. Environmental stress is the change of one or more
factors, such as: temperature, pH, presence of organic solvents, osmotic pressure, Mn2+ ions, blue
light or mutations (Volker, et al., 1995) (Gaidenko, et al., 2006). The function of stress receptors is
played by a multiprotein complex, called stressosome, which is formed in cytoplasm by RsbT, S, R,
RA, RB, RC, RD, and YtvA proteins. There is another protein kinase among these proteins – RsbT.
In unstressed cells RsbT is kept in stressosome in an inactive state. When stress occurs, this structure
is rearranged in order to release RsbT. Free kinase phosphorylates RsbS along with several RsbR
proteins prevent association of RsbT in the stressosome. Released kinase forms a complex with RsbU
phosphatase and activates it. Active RsbU dephosphorylates RsbV and general stress response is
turned on. Other proteins take part in stress sensing: ribosome building proteins (L11) and Obg, an
essential GTP-binding protein (Hecker, et al., 2007). These two proteins link stress response to
cellular translation and metabolic status. Following stress activation, over 150 proteins are
synthesized in the cell, using approximately half of the translational capacity of the cell, which allows
it to survive under stressful conditions. However, synthesis of these proteins is transient, reaching
top level after about 20 minutes after the change and decreases to the pre-change level in the next
Protein phosphorylation in bacteria. 2. General stress response – How to manage stress?
by Michał Obuchowski
15
dozen of minutes. The responsibility for quenching the stress response is assumed by phosphatase
RsbX protein, which dephosphorylates p-RsbS and p-RsbR proteins. Energy stress response depends
on activity of RsbP protein phosphatase, acting together with RsbQ α/β hydrolase. This pair of
proteins initiate dephosphorylation of p-RsbV in response to glucose, phosphate, or oxygen
starvation, presence of azide, or NO. Precise role of RsbQ in activation of phosphatase activity of
RsbP is not clear yet (Kaneko, et al., 2005). Summing up, in B. subtilis general stress response
pathway involves two protein kinases, RsbW and RsbT, together with three protein phsophatases,
RsbU, RsbP, and RsbX. The first protein kinase, RsbW, exhibits an inhibitory effect, while the second
activates induction of stress response. Similar situation can be seen in case of phsophatases – RsbU
and RsbP activate, while RsbX inhibits activation of SigB protein. Our knowledge of this matter is not
complete, because RsbU- and RsbP-independent activation of SigB pathway has been described
(Brigulla, et al., 2003). Similarity searches revealed that so far regulation of stress response is most
complex in B. subtilis, in comparison to other Gram-positive bacteria. Proper and accurate stress
response increases the rate of survival by about three orders of magnitude in case of environmental
stress, and by one in case of energy stress. Presence of one type of stress results in an increase of
stress tolerance in all possible conditions. Interestingly, mutants unable to respond to stress grow
normally in optimal, laboratory conditions, in contrast to mutants defective in rsbX gene, which grow
very poorly.
16
1TProtein phosphorylation in bacteria. 2. General stress response – How to manage stress?
by Michał Obuchowski
3. Control of nitrogen assimilation
Assimilation of nitrogen, one of the most important elements for life, is crucial in maintaining
the fastest growth rate. Preferred form of nitrogen is ammonia. NH4+ is assimilated into two amino
acids, glutamate and glutamine. These two amino acids are used as a source of nitrogen for other
nitrogen-containing biomolecules. This is the reason why one or both of these amino acids are
present in most cells, at concentrations higher than other amino acids. This concentration is
regulated not only in response to cell’s nitrogen requirements, but also in order to maintain
an osmotic balance between cytosol and the external medium. The biosynthetic pathways of
glutamate and glutamine are simple in comparison to importance of these compounds. In practice
there are two reactions: firstly, ammonia is assimilated to glutamate, which results in obtaining
glutamine; secondly, α-ketoglutarate and glutamine are converted into two molecules of glutamate.
Obviously, the most important enzyme for nitrogen assimilation is glutamine synthetase (GS). When
ammonia is present in excess, only a low level of this enzyme is present in the cells. However, when
environmental concentration of ammonia is a growth-limiting factor, the level of synthesis of GS is
elevated (Reitzer, et al., 1985). Changing availability of nitrogen influences expression of several
operons, which form Ntr regulon. This regulon is controlled by a two-component system, consisting
of NtrB (GlnL, NRII), receptor histidine kinase, and NtrC (GlnG, NRI), a response regulator.
In order to make this situation as clear as possible, we must go back to glutamine synthetase.
This enzyme consists of 12 subunits, organized in hexa-element rings. Each of the enzyme's subunits
is independently regulated by covalent modification (adenylation) and competitive allosteric control
by 9 small molecules (carbamyl phosphate, tryptophan, glycine, alanine, histidine, serine, AMP, CTP,
and glucosamine-6-phosphate) (Eisenberg, et al., 2000). Each molecule alone exhibits only partial
inhibitory effect, but the effects of multiple inhibitors are more than additive. Covalent modification
by adenylation has no direct effect on enzymatic activity of glutamine synthetase, but modified
subunits are more sensitive to allosetericall control (Nelson, et al., 2008). Concentration of particular
small molecules depends directly on the status of the cell, e.g. lack of amino acids may reflect
conditions of amino acid starvation. Covalent modification by reversible adenylation is catalysed by
adenyltransferase (ATase), a product of glnE gene (Rhee, et al., 1985). ATase activity is regulated in
two ways: alloseterically, by α-ketoglutarate and glutamine, as well as by interaction with two
proteins, PII (product of glnB) and GlnK, which are encoded by genes of the same names. While
presence of α-ketoglutarate inhibits adenylation of GS by ATase, glutamine has an opposite effect.
Control of ATase activity by PII and GlnK proteins is more complex. In vivo, these proteins form a set
of trimers: PII3, PII2GlnK, PIIGlnK2 or GlnK3. Each homo- or heterotrimer exerts a slightly different
modulatory effect on ATase activity (van Heeswijk, et al., 2000). But it is not the end. PII and GlnK
proteins are covalently modified by uridylyltransferase/uridylyl-removing enzyme (UTase/UR), which
is encoded by glnD gene (Adler, et al., 1975). Uridylation status of PII and GlnK regulates the effect of
its interaction with ATase. Unmodified homo or heterotrimer of PII and GlnK positively regulates its
activity, resulting in downregulation of GS. Uridinylated PII or GlnK exhibits an opposite effect.
But new question arise: how is UTase/UR regulated? Uridylyl-removing activity of this enzyme is
regulated by the presence of glutamine, in combination with Mn2+. Opposite activity is stimulated by
ATP and α-ketoglutarate. It is worth of noticing that listed molecules do not compete with each
other. Taking all this data into account, it can be concluded that UTase/UR and ATase are involved in
sensing intracellular concentration of glutamine. In other words, modulation of its activity reflects
Protein phosphorylation in bacteria. 3. Control of nitrogen assimilation
by Michał Obuchowski
17
cellular nitrogen status. Well, what does it have to do with protein phosphorylation? Transcription of
GS originates at two promoters: glnAp1 and glnAp2. The first one is a weak transcription promoter
recognized by RNA polymerase in complex with σ70 subunit. This transcription maintains basal level
of GS synthetase. The second is a strong promoter recognized by σ54 subunit, but as all
σ54-dependent promoters, it absolutely requires the presence of specific activator protein – in our
case NtrC. Unmodified NtrC protein forms a dimer, which binds DNA with low affinity and has no
effect on transcription. However, upon phosphorylation, NtrC protein forms a tetramer and binds to
the region placed 110 and 140 bases upstream of glnAp2 promoter with high affinity (Ninfa, et al.,
1987). This represses transcription from glnAp1 promoter, because NtrC binding sites overlap with
-35 and +1 regions. Phosphorylated NtrC exhibits ATPase activity, which is necessary for
rearrangement of closed complex of RNA polymerase sitting on plnAp2 promoter into an open
complex, which results in initiation of transcription. NtrC-binding motifs are placed quite far from the
promoter region. Activator contacts RNA polymerase as a result of simultaneous action of NtrC
protein and other DNA-bending proteins. This resembles action of eukaryotic enhancers of
transcription (Carmona, et al., 1997). This is one thing, but how does NtrC become phosphorylated?
The answer is simple: by its cognate histidine kinase, NtrB protein. Dimers of NtrB kinase become
phosphorylated at histidine residue in position 139 in a trans-autophosphorylation reaction. Then,
the phosphate group is transmitted to residue 54 on NtrC protein. Sounds simple? Not exactly.
Kinase activity of NtrB should be regulated, and it is. Kinase exhibits two opposite activities: protein
kinase activity described above, and protein phosphatase, specific for phosphorylated NtrC.
Phosphatase activity of NtrB depends on the interaction of this protein with PII and GlnK proteins
(Arcondeguy, et al., 2001). When nitrogen is not a limiting factor, both PII and GlnK are not
uridinylated. In this state, interaction with NtrB, which becomes a protein phosphatase, results in
rapid dephosphorylation of NtrC, thus leading to a decrease of expression of glutamine synthetase.
When cells require more glutamine, PII and GlnK become uridylylated, which prevents interaction
with NtrB. NtrB becomes an active kinase, leading to a rise in the concentration of phosphorylated
NtrC. As expected, in cells lacking NtrC kinase, expression of GS is always low. Unexpectedly, the level
of transcription of glutamine synthetase in cells lacking NtrB protein, which grow on glucose is
absolutely normal. How is it possible? The response regulator, NtrC, can be phosphorylated directly
by acetyl phosphate, which is produced at the end of glycolysis pathway (glucose→pyruvate→acetylCoA→acetyl phosphate). This way of phosphate flow functions as a feedback mechanism between
carbon and nitrogen metabolism in the cell.
The above description of regulation of nitrogen assimilation is very brief and lacks details of
many connections with other metabolic processes, which take place in bacterial cells. However,
this description sheds some light on its complexity. Global regulation of nitrogen metabolism
discovered so far is complex and subtle, yet able to adopt cell metabolism to large and rapid
fluctuations in nitrogen availability in the environment.
18
1TProtein phosphorylation in bacteria. 3. Control of nitrogen assimilation
by Michał Obuchowski
4. Many stimuli for one response of Vibrio cholerae
A Gram-negative bacterium, Vibrio cholerae is an agent of a threatening disease – cholera.
Currently people alerted by different media are aware of flu epidemic or anthrax, and only a very
limited number of men know, how deadly cholera may be. Flu virus or Bacillus anthracis (agent of
anthrax) needs a few days to cause death. Cholera is much faster – it takes only several hours to kill
a healthy person. Moreover, cholera has a long and well-proven historical record. The region of
Ganges river is a natural reservoir of V. cholerae, as well as coastal areas around the world. This
disease has been known in that region since recording the first pieces of written history. Cholera
came to Europe for the first time in 1817, along with establishment of modern trade routes (Childers,
et al., 2007). After that desease started to spread across the world. Since then there have been seven
recorded pandemics of cholera: 1817, 1830, 1852, 1870, 1899, and the last, which begun in 1961.
Cholera is characterized by voluminous watery diarrhoea (characteristic “rice water”), which leads to
rapid dehydration, hypovolemic shock, acidosis and death, if no appropriate treatment is employed.
If this disease be left untreated, mortality reaches 60%, but treatment can reduce this rate to about
1%. Cholera is transmitted in poisoned food or water. A healthy person requires relatively high
amount of inoculum of bacteria (about 106–1011), but malnourished or immunocompromised
individuals are much more sensitive (Sack, et al., 1998). Bacteria ingested orally pass through the
stomach and colonize the surface of intestinal epithelial cells. Next, the production of cholera toxin is
started. The toxin is endocytosed by epithelial cells. Then, it is moved into endoplasmatic reticulum
using retrograde transport, and finally transported into cytoplasm. Then, the toxin activates cellular
adenylate cyclase to rapid overproduction of cAMP. A high level of this nucleotide allows for massive
secretion of chloride anions into intestinal lumen, followed by large amounts of water (Field, et al.,
1965). Cholera attacks suddenly, because toxin production is precisely regulated and coordinated in
bacteria present on the surface of epithelial cells. How do bacteria do this? The story starts with ToxR
protein. ToxR is an unusual transcription activator, which shows homology to response regulator of
two component systems, but there are several differences. Firstly, ToxR protein is bound to the inner
cell membrane. The N-terminus is in the cytoplasm, whereas the C-terminus is in the periplasm. ToxR
requires presence of another protein, ToxS. ToxS is also a membrane-bound protein, but it has only
one domain in the periplasm. The precise role of ToxS is still unknown, but it is postulated that
stability and dimerisation of ToxR depends on ToxS (Matson, et al., 2007). ToxR-ToxS complex
influences expression of two porins: OmpU and OmpT. ToxR-ToxS complex activates expression of
the first of them, while it represses expression of the second. Balance between OmpU and OpmT is
essential for resistance to bile acids and intestinal colonisation. A major function of ToxR is activating
transcription of ToxT. However, ToxR, even together with ToxS, is unable to activate its transcription.
Achieving this requires the presence of another transcription activator, TcpP. TcpP with TcpH form
another pair of membrane-linked proteins required for virulence expression by V. cholerae. The role
of TcpH is to inhibit YeaL protease from degradation of TcpP (Beck, et al., 2004). Coordinated action
of ToxRS and TcpPH leads to activation of transcription of ToxT. This protein is a major activator
protein responsible for expression of ctxAB genes, coding for cholera toxin and other virulence
genes. It is simple, yet incomplete. When expression of toxT is turned on, it means that cells received
several different stimuli from the environment and cell density. How are these stimuli received and
integrated into regulation of expression of ToxT? Let us go back to the pair of previously described
proteins: ToxRS and TcpPH. The first pair is expressed constitutively, but the second is not.
Protein phosphorylation in bacteria. 4. Many stimuli for one response of Vibrio cholerae
by Michał Obuchowski
19
Expression of TcpPH is regulated indirectly by quorum sensing mechanisms. Let us see how it
happens. V. cholerae has two sensor kinases, CqsS and LuxQ, which respond to different autoinducer
molecules. Phosphate is passed to LuxU protein, and finally to LuxO response regulator (Miller, et al.,
2002). When cell density is low, both kinases phosphorylate LuxU protein, which in turn activates
transcription of four regulatory RNAs, called Qrr1–4 (Lenz, et al., 2004). The sRNAs interact with Hfq
protein in order to destabilize hapR mRNA. In absence of HapR protein, aphA gene is transcribed,
which leads to activation of expression of TcpPH protein. An opposite situation occurs at high
densities. When CqsS and LuxQ kinases sense high concentration of appropriate autoinducers, they
switch to phosphatase activity in response. This leads to dephosphorylation of LuxU, which transfers
the phosphate group to LuxO (Freeman, et al., 1999). Expression of hapR gene occurs and in turn the
synthesized HapR protein blocks transcription of aphA gene. AphA protein is required for activation
of transcription tcpPH, and together with ToxR regulon, the lack of it blocks transcription of cholera
toxin. In addition the activity of LuxO transcription activator is regulated by another two-component
system, VarSA. The VarAS system controls expression of three small RNAs, which inhibit the activity
of CsrA protein, a global regulator, able to activate LuxO protein independently of LuxU (Miller, et al.,
2002). Expression of virulence by V. cholerae is also coupled with biofilm formation, regulated in turn
by at least next two two-component systems. The VieSAB two-component system consist of VieS
(sensor kinase) and two response regulators, VieA and VieB. VieA contains domains typical for
response regulators: a phosphorylated recipient domain and a DNA-binding domain, while VieB
contains only a single recipient domain. VieA protein influences expression of about 400 genes, by
using the second messenger, a 3’,5’-cyclic diguanylic acid (c-di-GMP). Intracellular level of c-di-GMP is
regulated by its synthesis and hydrolysis ratio balance. VieA acts as a phosphodiesterase and
degrades c-di-GMP, which leads to down-regulation of biofilm formation and up-regulation of
virulence expression (Tischler, et al., 2005). The second two-component system, which regulates
biofilm formation by V. cholerae, is not fully characterized yet. So far we know that response
regulators, VpsR and VpsT, activate expression of operons encoding genes necessary for
exopolysaccharide production, which are in turn crucial for biofilm formation. Unfortunately its
cognate kinase has not been identified yet. It is worth of pointing out that HapR, a previously
described quorum sensing transcriptional regulator, down-regulates biofilm formation. The last
factor, which will be discussed in terms of its influence on virulence expression, is glucose. Glucose
level is reflected in the cell by concentration of cAMP, a cofactor of CRP protein. Binding sites of CRP
and AphAB proteins overlap at tcpPH promoter. It has been shown that CRP binding represses tcpPH
transcription, acting as a negative regulator of cholera toxin expression (Kovacikova, et al., 2001).
Summing up, regulation of virulence expression by cholera agent is complex. It depends on
numerous environmental and intracellular signals, such as temperature, osmolarity, pH, glucose
accessibility and quorum sensing. Complexity of this regulation and its linkage to other regulatory
pathways is necessary for successful colonisation of both the environmental niche and hosts.
20
1TProtein phosphorylation in bacteria. 4. Many stimuli for one response of Vibrio cholerae
by Michał Obuchowski
5. A very important question – to sporulate or not to sporulate?
Endospore formation processes triggered by miltistymulus, such as limitation of nutrients
and cell density by Bacillus subtilis, is one of adaptive responses used by Gram-positive bacteria. As a
result of this developmental pathway, a spore is produced. It is a dormant cell, highly resistant to a
variety of harsh conditions, which can maintain its potential for vegetative growth for extremely long
periods of time. Complexity of this process and its similarity to development of higher organisms
drew attention of many scientists in past decades. The existence of spores was for first time reported
simultaneously by Koch and Cohen in 1876. Currently, after nearly one and a half century of
research, our knowledge about the sporulation process is reasonable, yet still incomplete. In 1992
Losick and Stragier proposed the present model of molecular events during initiation of sporulation
(Losick, et al., 1992). A lot of information has been added since then, but the general view remains
unchanged. In the following sections you will find a brief description of the current state of
knowledge about spore formation in B. subtilis.
Vegetative form of B. subtilis can be found nearly in all niches across the world. This nonpatogenic, rod shaped, free-living bacterium exists inside humans and animals, on growing plants, in
soil, as well as in water. Growth in so numerous and varied environments is possible due to its ability
to form an almost indestructible structure – the spore. In response to nutrient limitation B. subtilis
can enter a sporulation pathway, which involves about 500 genes (1/8 of all genes present in this
bacterium) and lasts 6 - 8 hours (Ellermeier, et al., 2006). At this point we have to emphasize the
words “can enter”. Sometimes it is hard to realize that the sporulation pathway is the last hope for
bacteria, and there are several reasons why it is taken as a last resort. Sporulation is a long process –
8 hours for bacteria would translate into decades for humans. It is energy consuming – it must begin
before the energy source is exhausted. Finally, it becomes irreversible at approximately 1/3 of the
whole process. Before entering the sporulation pathway a cell “needs to analyze” all available
information and scenarios. Speaking in molecular terms, the regulatory circuit needs to be more
complicated than just a simple two-component system. And it is.
So far five histidine kinases have been identified, which “pump” phosphate into the
phosphorelay. These proteins are called KinA-E (Stephenson, et al., 2002). These kinases undergo
different expression and respond to different stimuli, e.g. KinB and KinC are autophosphorylated in
response to stimuli indicating high population density (Burbulys, et al., 1991). Next, phosphate is
transferred to Spo0F protein, which is the primary response regulator. The postulated role of Spo0F
is to integrate the phosphate flow from all kinases, but this protein does not directly influence gene
expression. The phosphate group is transferred from Spo0F to Spo0A by a phosphotransferase,
Spo0B. The effector protein of phosphorelay is Spo0A, which positively or negatively regulates
transcription of at least 40 and 80 genes, respectively, in a direct way. Indirectly, it influences close to
a few hundred of genes (Molle, et al., 2003). There are two classes of genes among the genes
activated by phosphorylated Spo0A (pSpo0A): genes of the first class are activated by a low
concentration of pSpo0A, while genes of the second remain unaffected, until concentration of the
phosphorylated form of Spo0A reaches a high level. One of the most important genes activated by
a low level of pSpo0A is abrB gene, coding for a repressor of many genes transcribed in the stationary
phase (Strauch, 1993). As was mentioned above, sporulation occurs in response to nutrient
limitation. CodY protein plays a major role in this process. This protein is a repressor of early
sporulation genes. CodY requires formation of a complex with GTP for proper functioning. Until the
Protein phosphorylation in bacteria. 5. A very important question – to sporulate or not to sporulate?
by Michał Obuchowski
21
cell has a sufficient level of nutrient, the intracellular level of GTP remains high and CodY-GTP
represses sporulation genes. When nutrients become a limiting factor, a drop in GTP concentration
inactivates CodY and transcription can occur (Serror, et al., 1996). It is worth of noticing that the
majority of genes repressed by AbrB or CodY are not exactly genes connected with sporulation.
They encode a variety of proteins expressed in order to help cells in adapting to stationary phase
conditions. The initial amount of phosphate transferred to Spo0F protein by histidine kinase is not
equal to that received by Spo0B, because phosphorylated form of Spo0F (pSpo0F) is a substrate for
three aspartyl phosphatases: RapA, RapB, and RapE (Perego, et al., 2001). Extensive
dephosphorylation of pSpo0F prevents entering sporulation pathway too rapidly. In response to
different physiological states of the cell, expression of particular phosphatases is induced, which
counteracts sporulation initiation (Perego, et al., 1994). Interestingly, the activity of Rap
phosphatases is carefully regulated by specific peptides, called Phr (their names also include
a corresponding character, i.e. PhrA inhibits RapA, etc.). Genes encoding phr peptides are often
located in the operon with target phosphatase. The product of translation of phr mRNA is a peptide
of about 40 amino acids in length. It has a signal peptide at N-terminus, which is responsible for its
translocation across the cell membrane. After transport occurs, signal peptidase cleaves the Phr
peptide to the length of about 14 amino acids. Next, it is processed on the extracellular side of the
membrane, to its final form of a pentapeptide. Then, the active form of Phr peptide is internalized by
an oligopeptide transport system, Opp (Rudner, et al., 1991). In the past, it was believed that Phr
peptides are secreted to the environment, and their internalization is somehow connected with cell
density. But should this assumption be correct, Phr would never reach sufficient concentration to
efficiently inhibit Rap phosphatases, because the enzymes and their peptide inhibitor are synthesized
in nearly equimolar ratio (Core, et al., 2001). Current interpretation of this phenomenon is that
export, processing and import of Phr peptides constitutes a system delaying entry into sporulation
pathway (Perego, et al., 2001). Let us go back to the cell. Decreased activity of RapA,B and E
phoshphatases allows for accumulation of pSpo0A at low level. This is sufficient for activation of
several very important genes: spo0A, kinA, and spo0H. The first gene does not require any
explanation. The second gene encodes the most efficient histidine kinase, of which activity pushes
the cell into the sporulation pathway. The last gene encodes an alternative sigma subunit of RNA
polymerase, sigmaH, which is activated during transition from logarithmic to stationary phase of
growth. Moving balance of transcription towards sigmaH dependent genes results in expression of
spoIIA and spoIIG operons, a well as spoIIE genes, along with other genes required for asymmetrical
cell division (Sonenshein, 2000). Among proteins encoded by operons spoIIA and spoIIG, are sigmaF
and sigmaE, respectively. They are two of the early sporulation-specific sigma factors. Both subunits
are kept in an inactive state by two different mechanisms, which will be described in the following
part. A cell possesses another checkpoint in the phosphorelay. It is a Spo0E phosphatase, together
with its two paralogs, YisI and YnzD. Expression of these phosphatases is independently regulated by
growth conditions, which favour vegetative growth (Perego, 2001). Their activity does not allow the
cell to initiate sporulation, if environmental conditions can still support growth. In addition to
information presented about the regulation of phosphate flow through phosphorelay, there are
several other regulatory circuits in the cell, which can delay initiation of sporulation, such as SinRSinI, Soj-Spo0J, KipI-KpA, Sda, Hpr, and probably several others, which have not been described yet.
SinR plays a similar role to AbrB and CodY — it represses genes involved in early sporulation and
biofilm formation. Its antagonist, SinI, is a sigmaH-dependent gene and it inactivates SinR by forming
22
1TProtein phosphorylation in bacteria. 5. A very important question – to sporulate or not to sporulate?
by Michał Obuchowski
a SinR-SinI complex. Soj and Spo0J proteins form a link between sporulation and replication. Until
completion of replication, free Soj inhibits transcription of spo0A and spoIIG. Upon completion of
replication ori regions are bound to cellular poles, Spo0J sequesters Soj at the poles, and Soj exhibits
its inhibitory action. KipI is an inhibitor of phosphate transfer from KinA to Spo0F. Its effect is
counteracted by KipA protein. Another protein, which influences activity of histidine kinase A, is Sda.
This protein inhibits KinA in response to DNA damage or block in DNA replication. Both regulatory
circuits, Sda and KipI-KipA, link sporulation initiation with DNA replication status and its quality. The
Hpr protein links sporulation with carbon availability and metabolism. If sufficient amount of carbon
is present, Hpr indirectly inhibits expression of oligopeptide permease, Opp, necessary for import of
Phr peptides and SinI, which abolishes activity of SinR. Unfortunately molecular details of Hpr
influence on Opp and SinI are unknown (Koide, et al., 1999).
When pSpo0A reaches a level sufficient for initiation of sporulation, a few things happen.
FtsZ moves from mid-cell to a polar location, in more less ¼ or ¾ of the cell length. The spoIIA and
spoIIG operons are extensively expressed, leading to synthesis of sigmaF and pro-sigmaE. Expression
of sigmaF is accompanied by two other important proteins: SpoIIAA and SpoIIAB. The SpoIIAB protein
is called an anti-sigma factor, because it forms a complex with sigmaF, which results in its
inactivation. The SpoIIAA protein is an anti-anti-sigma factor, and its role is binding SpoIIAB, which
leads to release of active sigmaF. SpoIIAB is a protein kinase. It is able to phosphorylate SpoIIAA on
Ser58, which leads to inactivation of SpoIIAA (Arabolaza, et al., 2003). Sporulating but predivisional
cells contain active sigmaA and sigmaH, inactive sigmaF (in complex with SpoIIAB), pro-sigmaE and
phosphorylated SpoIIAA (pSpoIIAA), along with a high level of pSpo0A. Prior to formation of septa,
chromosome must be also prepared for division. When Spo0A reaches level sufficient for initiation of
sporulation, replication of the chromosome is already completed and origin regions are moved to the
cell poles, which results in stretching the chromosome across the cell. Then, asymmetrical septum is
formed. Among many proteins necessary for septa formation, there is one worth looking at: SpoIIE.
This protein is crucial for activation of sigmaF in forespore compartment of the cell and it is also
involved in asymmetrical division. This protein has twelve transmembrane regions located in
N-terminal part, and a PPM phosphatase located in the C-terminal part (Adler, et al., 1997) (Arigoni,
et al., 1999). Closing septa catches about 30% of forespore-destinated chromosome in prespore
compartment. How can the cell distinguish between the mother cell and the forespore
compartment? How is sigmaF specifically activated only in the forespore compartment? The first step
in finding an answer to these questions is understanding the function of SpoIIE protein.
The N-terminal part of SpoIIE is submerged in the cellular membrane, and somehow this protein is
sequestered in the new septa, probably on both sides of it. The forespore compartment is much
smaller, so dephosphorylation of pSpoIIAA by SpoIIE should be relatively quick, in comparison to the
mother cell compartment. It seems possible, but this is not the complete answer. To be precise, we
need to focus on the SpoIIAB kinase and its mechanism of action. SpoIIAB exists in two
conformations: open and closed. In the first conformation, the nucleotide binding pocket is
accessible for cytoplasmatic ATP. Moreover, ADP generated during the phosphorylation event can
leave the enzyme. In closed conformation, nucleotides can not enter or escape from the pocket. The
cycle of SpoIIAB begins, when the enzyme in open conformation binds ATP. Then, the substrate
(SpoIIAA) can be bound. This event forces conformation of SpoIIAB-ATP to change from open to
closed. The result of this is a ternary complex: SpoIIABclosed-ATP-SpoIIAA. At this stage
phosphorylation occurs, and the complex consists of SpoIIABclosed-ADP-pSpoIIAA. Phosphorylated
Protein phosphorylation in bacteria. 5. A very important question – to sporulate or not to sporulate?
by Michał Obuchowski
23
form of SpoIIAA no longer stays bound to SpoIIAB, and is released. ADP nucleotide is bound to
the kinase and remains in closed conformation. Precisely at this point the crucial step in sigmaF
activation takes place. Conformational change from SpoIIABclosed-ADP to SpoIIABopen-ADP is slow.
Considering the situation in predivisional cell of mother cell compartment, the slow shift of
conformation of SpoIIAB from closed to open is not a problem, because activity of SpoIIE
phosphatase can be ignored. However, the situation is different in the forespore. pSpoIIAA is
extensively dephosphorylated by SpoIIE. This leads to an increased level of dephosphorylated
SpoIIAA, which is characterized by affinity to SpoIIAB. When SpoIIAA binds to SpoIIABclosed-ADP,
a ternary complex of SpoIIABclosed-ADP-SpoIIAA is formed, which blocks kinase activity of SpoIIAB,
because the nucleotide cannot be released. You need to remember that free SpoIIAB shows affinity
towards sigmaF. Before asymmetrical division, all sigmaF is sequestered in the SpoIIABopen-ATPsigmaF complex. After septa formation, activity of SpoIIE results in accumulation of unphosphorylated SpoIIAA able to bind SpoIIAB, which in turn leads to release of active sigmaF (Yudkin, et
al., 2005). It has also been postulated that decreased concentration of ATP in the forespore
compartment can affect the activity of SpoIIAB kinase, but this plays only a minor role in activation of
sigmaF. What does the situation look like now? The mother cell contains pSpo0A, SpoIIAB-sigmaF,
pSpoIIAA, and pro-sigmaE, while the forespore contains pSpo0A, SpoIIAB-SpoIIAA, and active sigmaF.
This is clear, but is active sigmaF the only factor responsible for differential gene expression in the
forespore? Of course not. The presence of phosphorylated Spo0A is counteracted by dephosphorylated SpoIIAA, which leads to a decrease of expression of genes activated by pSpo0A. In the
case of spoIIA operon, the situation is more interesting. As was said before, when asymmetric septa
are formed, only 30% of chromosome located near origin is trapped in the forespore compartment.
The spoIIA operon is located in the distal part of the chromosome and it is simply absent in the
forespore immediately after asymmetrical division. Because of this reason, the new molecules of
SpoIIAB cannot be synthesized, and activation of sigmaF is faster. After approximately additional
15 minutes the rest of the chromosome is translocated into the forespore due to activity of SpoIIIE
protein, but this delay is sufficient for decrease of Spo0A level, which prevents expression of spoIIA
and spoIIG operons (Sharp, et al., 2002). At the same time, in the mother cell compartment, SpoIIAA
remains phosphorylated and pSpo0A can activate gene expression. In this part of the cell the most
important element is spoIIG operon, encoding pro-sigmaE. Undisturbed activity of pSpo0A results in
an increase of concentration of pro-sigmaE in the mother cell compartment. At this point, you should
remember that pro-sigmaE was also synthesized before asymmetrical division, and a portion of it was
trapped in the forespore compartment. This portion is eliminated from the forespore by proteolytic
degradation. Stability of pro-sigmaE is the same in both parts of the cell, but in the forespore its
expression is blocked by the presence of SpoIIAA (Fujita, et al., 2002). The level of pro-sigmaE is
maintained in the mother cell, because pSpo0A continuously activates expression of spoIIG operon.
So now active sigmaF is present in the forespore, but sigmaE in the mother cell remains inactive.
How can the cell activate it? Presence of active sigmaF in the forespore allows for expression of
approximately 48 genes belonging to its regulon (Wang, et al., 2006). Among them is the spoIIR gene.
SpoIIR protein is secreted to intermembrane space between the mother cell and the forespore.
Postulated function of SpoIIR is transmitting of a signal from the forespore, which is required for
activation of sigmaE in the mother cell (Arabolaza, et al., 2003). SpoIIR interacts with membrane
bound protease, SpoIIGA, which is expressed in the mother cell under control of Spo0A and SinR.
Interestingly, membrane localisation of SpoIIGA depends on SpoIIE protein (Fawcett, et al., 1998).
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1TProtein phosphorylation in bacteria. 5. A very important question – to sporulate or not to sporulate?
by Michał Obuchowski
Activated SpoIIGA cleaves pro-sigmaE to sigmaE in the mother cell compartment. Why do sporulating
cells usually form only one forespore? In the mother cell compartment FtsZ ring still exists, i.e. it is a
potential site for another septum. Also there is a high level of pSpo0A, which activates expression of
all genes involved in asymmetrical division. Typically, activation of sigmaE in the mother cell
compartment prevents formation of another septum and the second forespore compartment. In
relatively rare situations a scenario is possible, where a single cell forms two forespore
compartments after two asymmetrical divisions (Eldar, et al., 2009). Summing up, sigmaF is activated
in the forespore compartment following asymmetrical division. At the same time, chromosomal DNA
is pumped into the forespore and septation is completed. Approximately at the same time activity of
sigmaF allows for activation of sigmaE in the mother cell compartment, which prevents another
septation and is necessary for proceeding to the next step of sporulation. The fact that activation of
sigmaE depends on transcription of sigmaF-dependent gene (spoIIR) seems to be a well designed
security system, which does not allow proceeding to the next step of sporulation without completion
of the previous step.
SigmaE regulon currently consists of 154 genes, but this number may increase, because new
genes are still being identified and described (Steil, et al., 2005). The major task of sigmaE-dependent
genes is to drive the engulfment process and after its completion to initiate synthesis of the spore
coat. Engulfment is a highly coordinated process of surrounding the prespore by the mother cell. This
process requires coordinated expression in the mother cell of such genes as spoIID, spoIIP,
and spoIIM, together with spoIIQ expression in the forespore. SpoIIP and SpoIID exhibit enzymatic
activity necessary to degrade peptydoglican of the cell wall and can drag mother cell's membrane
(Morlot, et al., 2010). At the end of the engulfment process the mother cell's membrane must be
fused. This requires the presence of SpoIIIE and SpoIIQ proteins (Sharp, et al., 2003). Successful
ending of engulfment is necessary for activation of late sporulation sigma factor, sigmaG, in
the forespore, and sigmaK in the mother cell. Gene expression, which occurs in the mother cell leads
to appearance of SpoIIID and GerR proteins, as well as pro-sigmaK. SpoIIID and GerR are primarily
repressors of sigmaE dependent genes (Eichenberger, et al., 2004). However, SpoIIID also activates
expression of several genes, including the one, which encodes pro-sigmaK. Other proteins, SpoIIIAH
and SpoIIIJ, are required for activation of sigmaG in the forespore. This protein interacts with SpoIIQ,
leading to activation of sigmaG in the forespore upon completion of engulfment. Unfortunately,
precise molecular mechanism of activation of sigmaG remains unknown. Expression of sigIIIG (sigG)
is driven by sigmaF in the forespore, but this subunit is synthesized as inactive precursor and requires
proteolytic activation. Appearance of active sigmaG in the forespore forces expression of its regulon,
which consists of 113 genes (Steil, et al., 2005). Activation of these genes results in condensation of
the chromosome in complex with SASP proteins, along with proteins preparing forespore for
germination, when it encounters nutrients (Wang, et al., 2006). Active sigmaG leads to expression of
spoVT gene, which encodes a transcription regulator. All genes belonging to the sigmaG regulon can
be subdivided into three classes. Genes belonging to the first of them require only presence of
sigmaG. Expression of genes of the second class is activated by sigmaG and repressed by SpoVT.
Genes of the third class require sigmaG and activation by SpoVT. The last class includes spoIVB, a
gene, of which expression in the forespore is necessary for proteolytic activation of pro-sigmaK in the
mother cell compartment. Let us go back to the mother cell compartment. Scanning of the
chromosome for sigmaK gene yields surprising results: the gene is interrupted by a 48-kb element
called skin (Stragier, et al., 1989). This element is probably a phage-related sequence, which encodes
Protein phosphorylation in bacteria. 5. A very important question – to sporulate or not to sporulate?
by Michał Obuchowski
25
about 60 genes. These genes include spoIVCA, product of which is a recombinase, cutting out the skin
element and reconstituting functional sigK gene by means of fusion of spoIIIC with spoIVCB.
Rearranged sigK gene is expressed under control of sigmaE and SpoIIID, producing pro-sigmaK.
SpoIVFB protein is responsible for the activation of pro-sigmaK to sigmaK. Its activity is modulated by
SpoIVFA and BofA (Cutting, et al., 1990) (Ricca, et al., 1992). 132 genes belonging to sigmaK regulon
are mainly responsible for coat formation and lysis of the mother cell (Steil, et al., 2005). Among
them is gerE gene, which encodes another transcription factor, which in turn acts as a repressor or as
an activator of various genes. By activating sigmaK-dependent regulon, the cell completes
developmental programme of spore formation. The spore is now complete and persists in the
environment for a very long time (Russell, et al., 2000).
When sigma factor cascade occurs, at least two important things happen: synthesis of the
cortex surrounding the forespore, which is placed between the membranes, and coat assembly on
the surface of the outer membrane. Let us first look at cortex synthesis. The cortex is basically
a peptidoglycan (PG), and it can be divided into two layers. The first layer is placed directly on the
outer surface of the inner membrane. It is called germ cell wall, and its structure is very similar to the
regular cell wall. This thin layer of peptidoglycan is synthesized before the rest of the cortex, due to
activity of the forespore, i.e. enzymes involved in its synthesis are expressed in forespore. Moreover,
necessary precursors of the germ cell wall are also synthesized in the forespore (McPherson, et al.,
2001). The second layer of the cortex is a modified peptidoglycan, which includes modified sugar,
muramic-δ-lactam devoid of the peptide chain (Popham, 2002). In this case there is a different
situation. Enzymes responsible for cortex synthesis (SpoVB, SpoVD, SpoVE) are expressed under
control of sigmaE in the mother cell compartment. They are placed in the outer membrane during
engulfment. However, synthesis of the cortex does not take place, since there are no available
precursors. After completion of engulfment and activation of sigmaK, the expression of the mur
genes takes place. This set of genes encodes enzymes, which synthesize precursors for cortex
polymerisation (Vasudevan, et al., 2007). Such organisation of cortex synthesis does not interfere
with transport between the forespore and the mother cell. When sigmaK becomes active, the
developmental programme of the forespore is nearly completed (sigmaG is already active), and the
whole preparation for dormancy state is finished. Synthesis of the cortex will lead to dehydratation
of the forespore and it has direct influence on resistance of the spore to environmental conditions.
In other words, forespore is ready for dormancy and completion of its shields depends mostly on the
mother cell compartment. If the cortex is synthesized too early, necessary preparations in the
forespore may stay uncompleted, because polymerisation of the cortex will lead to a too early
energy drop in the developing spore. Expression of both cortical synthetases under control of sigmaE
and precursor of peptidoglycan synthetase under control of sigmaK is a simple and elegant system,
assuring a proper start of PG synthesis, without any unnecessary delay.
Let us now concentrate on coat assembly. The spore coat is a wide layer of proteins, which
cover the spore. The number of various proteins, which compose the spore coat, is still unknown.
Currently over 50 of them have been identified, but it is extremely hard to predict how many have
not been characterized yet (Kim, et al., 2006). What is the cause of this situation? Proteomic analysis
became a routine, high-throughput experiments. Why does precise composition of the spore coat
remain unsolved? The answer is quite simple: proteins, which build the spore coat are often
corsslinked with rarely occurring chemical bonds, which makes them insoluble and difficult to
analyze. Genes, which encode coat proteins are expressed under control of mother cell-specific
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1TProtein phosphorylation in bacteria. 5. A very important question – to sporulate or not to sporulate?
by Michał Obuchowski
sigma factors. Let us start from the beginning. The first set of coat proteins is synthesized when
engulfment starts. After its completion the outer membrane is covered by SpoIVA protein, the first
coat protein, of which presence is absolutely required for proper assembly. But how can SpoIVA
recognize the outer membrane of the forespore and not polymerise on the membrane of the mother
cell? The answer is quite simple. The forespore expresses SpoVM, a membrane protein, which
interacts with SpoIVA and recruits it for polymerization on the membrane. Homopolymers of SpoIVA
form the base layer for further coat assembly (Ramamurthi, et al., 2008). At least 10 other proteins
are deposited on the SpoIVA layer, starting formation of the inner spore coat. Of course not all
proteins which build inner coat interact directly with SpoIVA. At least ten more proteins, which do
not interact with SpoIVA, have been identified in this part of the spore coat (Kim, et al., 2006).
When spores were analyzed by electron microscopy, it was found that the inner coat has lamellar
structure. Deposition of the outer coat is started by CotE protein, which forms the top layer of the
inner coat, similarly to SpoIVA (Takamatsu, et al., 2002). In the outer coat 27 protein have been
identified so far. Most of them somehow interact with CotE, but not all. It should be pointed out that
this layer contains the third morfogenic protein, CotH. In a very simplified model of interaction
network in the outer coat all proteins interact with at least one of CotE or CotH proteins. The outer
coat is an electron dense layer without visible structures under the electron microscope. Interactions
between coat proteins are much more complicated than outlined above, but all authors agree that
SpoIVA, CotE, and CotH play a major role in proper assembly of this protective layer of the spore.
The final event of the sporulation pathway is lysis of the mother cell and release of the spore
into the environment. As was mentioned at the beginning, sporulation is time- and
resource-consuming. It is the reason why cells try to find a way to delay it or even avoid it at the cost
of the rest of population. The reason for it may be simple, yet interesting — the decision to enter the
sporulation pathway is irreversible. If cells start sporulation, and environmental conditions improve,
these cells can not immediately return to vegetative growth, and if environment can no longer supply
nutrients for the cell, sporulation may be not completed, which makes the decision very risky.
When a spore is successfully released it is well protected form all natural environmental
conditions, which could lead to its death. But how to check if environment improves enough, so that
it can support vegetative growth again? The answer to this question is the fact that every spore is
equipped with several receptors sensing low molecular components, the presence of which should
suggest improved conditions. Mode of action of these receptors is different in different species
which can produce dormant spores. Usually germination is initiated by simultaneous presence of
a few different low molecular compounds, such as amino acids, nucleotides, sugars or other
compounds (Ross, et al., 2010). It is very important to realize that germination must be successful
at the first approach. During germination spores irreversibly release different ions and other
molecules. There is no evidence of active transport or metabolism of the germinant during
germination, which appears to be an essentially biophysical process (Moir, 2006). What do spores
produced by B. subtilis look like in terms of germination receptors? Up to date, three operons coding
for germination receptors have been characterized: gerA, gerB, and gerK. Each of these has three
genes, the products of which build the receptor (Paidhungat, et al., 2002). GerA receptor is
responsible for sensing the presence of L-alanine. GerB together with GerK sense the presence of
asparagines, glucose, fructose and potassium ions. When appropriate germinants are detected, the
dormancy state ends and the spore starts to germinate. The whole germination process can be
divided into two steps: in the first step the spore releases different cations (such as H+, Zn2+, K+, etc.),
Protein phosphorylation in bacteria. 5. A very important question – to sporulate or not to sporulate?
by Michał Obuchowski
27
followed by release of a complex of Ca2+ and pyridine-2,6-dicarbozylic acid (dipicolinic acid, DPA).
Deposit of DPA is high in dormant spores and reaches even 10% of dry weight of the spore (Setlow,
2003). Following the release of Ca2+-DPA, water fills the spore's space, which results in partial
rehydratation of the spore core. An already described movement of ions and water increases pH
inside the spore by more than one unit, which is sufficient for enzyme’s activation, when combined
with hydratation of the core (Jedrzejas, et al., 2001). The second step of germination starts when
cortex-degrading enzymes, SleB and ClwJ, become active. Degradation of the cortex is necessary for
further core hydratation and its expansion. After completion of hydratation of the spore core,
metabolic processes are started and germination is over — the coined term is “outgrowth”. This
stage starts when metabolism of spore is restarted. During this stage the SASP protein, which was
bound to the chromosome, is degraded, supporting germinating spore with amino acids. In addition,
the first new macromolecules are synthesized. Finally, new vegetative cells are released from
the spore coat.
Germination receptors are important for induction of germination in response to
the presence of low molecular compounds, but it is not the only way to break spore dormancy. It has
been reported that the product of peptidoglycan degradation can trigger germination, due to action
of protein kinase, PrkC, placed in the inner membrane of the spore (Shah, et al., 2008). In addition,
a few non-nutrient germinats have been characterized, such as high pressure, presence of Ca2+-DPA,
cationic surfactants, lysosyme or salts (Setlow, 2003).
Dormant spores are quite sensitive to the presence of germinants. Germination can be
sufficiently triggered, if necessary germinants are present for only a short time (minutes).
Non-nutrient germination is receptor-independent. However, its molecular mechanisms of action is
still unknown. Once the spore starts the germination process, it cannot be reversed. Current
understanding of ending spore dormancy is incomplete, but reasonable approximations can be
constructed using available information, which is derived mostly from studies of Bacillus subtilis.
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1TProtein phosphorylation in bacteria. 5. A very important question – to sporulate or not to sporulate?
by Michał Obuchowski
6. Bioluminescence – in a mass we will shine like stars
Several organisms of different taxa are able to produce light. These can be simple organisms,
such as bacteria, but also much more complicated, like dinoflagellates, insects, cephalopods or fish.
Of course it is not a common thing, but numerous examples suggest that in certain environments
luminescence can be useful. Many of you were probably able to see with your own eyes a firefly
during summer after dusk. Light produced by this insect is used to attract female fireflies. Some
octopus species use bioluminescent ink to repel predators and attract prey. For similar reasons light
is used by fish living deep in ocean. Light emission makes it easer to recognize each other or
to attract potential prey.
Why do bacteria emit light? These organisms are too small to be seen, light emitted by single
cell is too weak to be noticed by another inhabitant, and production of light obviously requires
a reasonable amount of energy and oxygen. Analysis of enzymatic apparatus necessary for light
production indicates that it is too complicated to be maintained without reason. Certainly, it was not
a single mutation which created enzymes capable of light production. Luciferins are a class of
pigments of biological origin, which can be oxidized by luciferase enzyme, producing oxoluciferins
and light. This is a common basis of bioluminescence. However, closer examination of different
bioluminescent organisms revealed that chemical structures of luciferins are quite different and it is
unlikely that they have a common ancestor. Let us look back at bacteria. Bacterial abilities to emit
bioluminescence have been known by humans for a very long time. Many tales exist, especially
between sailors, describing luminescent sea water surrounding moving ships. Usually every tale
contains a bit of truth, so do these. When a ship is in motion, it creates a water disturbance in front
of the hull and behind it. Especially in these two places water is aerated and microorganisms which
are present on the surface of water get more oxygen and can produce bioluminescene.
This phenomenon is rare, because it requires the presence of a mass of bacteria, which are capable
of light production. Recently, it has been discovered that bacteria do not produce light at low density
of cells. But how can they know if the density is sufficient? In order to decide this, microorganisms
have evolved a quorum sensing mechanism. In simple words, it is a system, which can provide
information about population density with specificity towards own and other species. This
mechanism was discovered over 40 years ago in a marine bacterium, Vibrio fisheri (Nealson, et al.,
1970). Later on, similar systems were found to be present in many other bacteria. Studies of different
systems of quorum sensing revealed that bacteria use different molecules as signals. These
molecules can be grouped into a few groups. Acyl homoserine lactones are mainly used by Gramnegative bacteria, while short peptides are used by Gram-positive bacteria. In addition, other signal
molecules have been detected, such as furanosyl borate diester, which is produced and detected by
Vibrio harveyi (Cao, et al., 1989).
Let us focus on Vibrio harveyi. It is a free-living marine bacterium, present in the Baltic Sea,
capable of being a pathogen (Austin, et al., 2006). What makes this particular microorganism so
interesting? The fact that it is a polyglot. If we consider signal molecules to be languages, V. harveyi is
able to “speak” at least in three languages. How is it organized? The first language, based on acyl
homoserine lactones, is common among Gram-negative bacteria. Chromosome of V. harveyi encodes
the luxLM protein, of which product synthesizes 4-hydroxyl-C4-homo-serine lactone (Cao, et al.,
1989), which will be called AI-1. These molecules diffuse freely into the environment. Their presence
can be sensed by any bacteria containing an appropriate receptor. In case of our bacteria, the
Protein phosphorylation in bacteria. 6. Bioluminescence – in a mass we will shine like stars
by Michał Obuchowski
29
presence of AI-1 is registered by LuxN protein, a membrane-linked histidine receptor kinase (Bassler,
et al., 1993). LuxN kinase contains a periplasmatic receptor domain and a cytoplasmatic kinase
domain with a conserved histidine residue. When AI-1 molecule is absent, kinase undergoes
autophosphorylation and phosphate is transferred to LuxU phosphotransferase, and in turn to LuxO
response regulator. LuxO protein has typical composition and consists of a recipient domain and a
DNA-binding domain. Phosphorylated LuxO, along with σ54 subunit, activate expression of five small
regulatory RNAs. These sRNAs work in concert with Hfq chaperone to decrease the stability of luxR
transcript, thus blocking activation of expression of genes required for bioluminescence (Tu, et al.,
2007). The LuxR protein is a master regulator of expression of genes involved in quorum sensing. Up
to date, we have only few bits of information about genes belonging to this regulon, but it is already
known that this protein influences bioluminescence, production of exopolysaccharide, siderophore,
expression and secretion of metalloproteases and probably some virulence genes (Milton, 2006).
That is first ‘language’. Now let us proceed to another.
The second Vibrio harveyi signalling molecule is 3A-methyl-5,6-dihydro-furo(2,3D)(1,3,2)dioxaborole-2,2,6,6A-tetraol, called simply AI-2 (Surette, et al., 1999). AI-2 is synthesized by
LuxS and as was mentioned before, it is released into the environment. In order to detect it,
V. harveyi encodes the second membrane-linked histidine kinase, LuxQ. But this kinase is not able to
directly sense the presence of AI-2. It requires the presence of periplasmatic LuxP protein. It is
believed that LuxQ recognizes LuxP-AI-2 complex. Activity of LuxQ kinase is regulated in the same
manner as LuxN, i.e. when AI-2 is absent, kinase is active and “pumps” the phosphate group to LuxU
phosphotransferase. In other words, LuxQ and LuxN work synergistically. Presence of two
independently regulated kinases in such a system allows the cell to distinguish four different states:
no autoinducers, only AI1, only AI-2 and both compounds together. However, obtained results
suggest that this pathway acts like coincidence detector and both signal molecules need to be
present for full activation of bioluminescence (Mok, et al., 2003). Presence of only one autoinducer in
the environment has only a minor effect on bioluminescence. Due to simultaneous detection of AI-1
and AI-2, V. harveyi can distinguish between situations in which it is in a mono-culture or in a multispecies consortium.
To make the situation little more complicated, in 2004 another two-components system
involved in V. harveyi quorum sensing was described (Henke, et al., 2004). This third system consists
of CqsA, which produces signal molecule termed CAI-1 (cholera autoinducer-1), chemical structure of
which remains unknown. Presence of CAI-1 is sensed by membrane-linked histidine kinase CqsS,
which as was previously described, transfers phosphate group to LuxO via LuxU. As was mentioned
above, all three kinases are active when the signalling molecules is absent. When a proper
autoinducer appears, kinase activity is switched to phosphatase and the flow of phosphate is
reversed. It allows for a quick change of phosphorylation status of LuxO protein. The described
system looks quite complicated, but certainly there are gaps in our knowledge. We do not know why
different genes controlling quorum sensing are regulated differentially in response to different
autoinducers (Waters, et al., 2007). Why is it so important? In certain conditions V. harveyi causes
reasonable losses in aquacultures. Due to increased multi-drug resistance, there is an increased need
of searching for alternative methods of controlling it. Currently there are several reports available,
describing V. harveyi with impairment of one or more quorum sensing signalling pathways, which
show different pathogenicity towards different invertebrates or vertebrates. These observations give
new hope of finding new ways to control these bacteria (Defoirdt, et al., 2008).
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1TProtein phosphorylation in bacteria. 6. Bioluminescence – in a mass we will shine like stars
by Michał Obuchowski
Finally let’s go back to the question why do bacteria produce light. When bacteria live under
a certain layer of water, they are protected from DNA damage induced by direct UV light.
Microorganisms can actively swim in order to control their depth, but cannot resist massive motions
of water, e.g. during a storm. Exposure to elevated levels of UV light causes increased damage of
DNA. Usually bacteria can repair some of these errors using photoreactivation pathways, but
if bacteria are exposed to light and after that quickly pushed back into deep water, the light is
insufficient. So, marine microorganisms compensate for the lack of sunlight by bioluminescence.
If population density is low, production of light is very inefficient in terms of DNA repair. However,
at high density bioluminescence becomes more intense and sufficient for photoreactivation reactions
(Czyż, et al., 2003).
Protein phosphorylation in bacteria. 6. Bioluminescence – in a mass we will shine like stars
by Michał Obuchowski
31
7. Genetic competence – to be competent or not?
The ability to uptake free DNA from environment has been detected in both Gram-positive
(Bacillus, Streptococcus) and Gram-negative (Acinetobacter, Campylobacter, Haemophilus,
Helicobacter, Neisseria, Vibrio) bacteria (Lorenz, et al., 1994). This unique mechanism of enrichment
of the chromosomal content may help bacteria survive in harsh conditions. When available nutrients
become insufficient for bacteria growing exponentially, they become motile, which enables them to
actively search for a new source of nutrients. If the state of nutrient deprivation continues, cells
enter the stationary phase and secrete a set of degrading enzymes in order to mobilize nutrients
from alternative sources. In parallel, starved cells start to produce a variety of antimicrobial
compounds in order to rip off competitors. If all these actions do not improve the nutrient status,
cells become competent and finally start forming spores (Hamoen, et al., 2003). Bacteria are usually
able to take up DNA of any origin (phage, plasmid, chromosomal) without any sequence specificity.
However, in case of Neisseria or Haemphilus specific sequence is required.
In Bacillus subtilis the master regulator of competence regulon is ComK protein.
This regulator positively regulates its own expression together with approximately hundred genes
involved in this process (Kovacs, et al., 2009). The expression of comK is subject to a sophisticated
regulatory system on transcriptional and posttranscriptional level. Let us discuss it step by step.
Transcription of comK is positively regulated by its own product, the ComK protein, together with
DegU protein. Negative control is conducted by Roc, AbrB, and CodY proteins (Hamoen, et al., 2003).
ComK recognizes a specific sequence, called AT-box, and binds to the minor groove of DNA. These
boxes are separated by a certain distance, which distinguishes them in terms of ComK binding into
high-affinity and low-affinity. In case of comK, AT-boxes create a low-affinity binding site for ComK.
When it is in the cell at a low concentration, it is unable to bind and activate its own expression. In
such conditions an auxiliary activator is necessary – the DegU protein (Hamoen, et al., 2000). The
DegU protein is a response regulator form of the DegS/DegU two component regulatory system,
which controls production and secretion of degrading enzymes in response to nutrient deprivation
(Kunst, et al., 1988). DegS is a membrane-linked histidine kinase, which phosphorylates DegU
response regulator in response to unknown stimuli. DegU protein activates expression and secretion
of degrading enzymes in phosphorylated state. Phosphorylated form of DegU has no affinity towards
its site, which is located between AT-boxes at comK promoter. However, unphosphorylated DegU
binds to its site and, due to interaction with ComK, enhances its binding capacity to its own site. This
is a biological version of the ‘OR’ logical gate. Cells can express degrading enzymes or become
genetically competent. When ComK reaches high concentration, the presence of DegU is no longer
needed. While that is a mechanism of activation of comK transcription, let us look at inhibition.
The two proteins AbrB and CodY were mentioned earlier, when we spoke about sporulation.
In case of competence their role is similar. Presence of AbrB does not allow entrance into
competence during vegetative growth and CodY links energy status of the cell with its developmental
pathway. The third negative regulator of comK expression is called Rok (Hoa, et al., 2002).
This protein binds directly to the comK promoter and represses it. In addition, Rok represses
expression of its own promoter. When ComK appears in the cell, it can bind specifically to rok
promoter and down-regulate it. Release of Rok repression allows for a rapid increase of ComK level
in the cell, which enables the cell to enter the competence pathway. Regulation of transcription of
comK, as you have probably already noticed, is much more complicated. The level of AbrB protein
32
1TProtein phosphorylation in bacteria. 7. Genetic competence – to be competent or not?
by Michał Obuchowski
depends on the level of phosphorylated Spo0A, and this depends on functioning of
the phosphorelay. It has been postulated that additional regulators, such as Med or SinR, act at comK
promoter as well, but their role in developing competence is not clear yet. As was mentioned at
the beginning, expression of comK is also extensively regulated at post-transcriptional level by
regulation of its proteolysis rate. ComK is degraded by ClpC-ClpP complex, but ClpC exhibits no
affinity for this protein. An additional protein, MecA, is necessary for efficient degradation of ComK
by ClpCP. On one side this protein exhibits affinity for ComK and on the other side the affinity for
ClpCP complex. It was shown that MecA recognizes FMLYPK motif placed at C-terminus of ComK
(Prepiak, et al., 2007). MecA forms the ComK-MecA-ClpC complex, in which ComK is rapidly
degraded. In other words, MecA recruits ComK to degradation by ClpCP. But new question arises: the
expression of MecA or ClpC is relatively constant, so how can this degradation pathway be
a regulatory system? The question remained unsolved until it was observed that at high cell densities
ComK was degraded in a slower manner.
It was already known, that competence somehow depends on quorum sensing machinery.
Another two component system is present here: ComP/ComA, which is involved in competence.
ComP is another membrane-linked histidine kinase, but in this case the ligand is known – it is the
ComX peptide. This signal peptide is synthesized as the end-product of comX gene expression. ComX
is expressed as 55 amino acid long peptide but it is inactive. ComQ protein is required for
its processing and secretion. When the final form of ComX is released into the environment, it is 9–10
amino acid long (Ansaldi, et al., 2002). Upon binding of ComX, ComP becomes active and
phosphorylates ComA response regulator. But presence of ComX is insufficient for triggering
competence. Phosphorylated ComA is a substrate of RapC phosphatase. In order to overcome this
block, PhrC peptide must inhibit RapC phosphatase activity. It is postulated that PhrC is another
quorum sensing peptide, which is internalized from environment by Spo0K permease (Hamoen, et
al., 2003). Accumulation of phosphorylated ComA activates expression of srf operone, which encodes
four very large enzymes responsible for surfactin production. Among these four large genes (this
operon is 36kb long) is hidden the fifth gene, comS. In contrast to surfactin synthetase enzymes, this
one is only 46 amino acid long, but its role is important. An increasing level of ComS allows it to
compete with ComK for MecA. This competition decreases the rate of proteolysis of ComK and allows
for developing competence. But when the ComS-MecA-ClpCP complex is formed, ComS is not the
only degraded enzyme. This complex also stimulates degradation of MecA protein by ClpCP, which
results in a decrease of its concentration and additional increase of half-life of ComK (Ogura, et al.,
1999). Co-expression of srf and comS is an example of smart and economical regulation. Increased
level of ComS favours development of competence, but at the same time the cell starts to synthesize
and secrete surfactin. This powerful biosurfactant provokes lysis of surrounding bacteria and this
increases the chance of finding DNA, which can be uptaken. When all the necessary modulators of
intracellular level of ComK point at competence, its concentration increases very rapidly. However,
when all conditions appear to be optimal, only 10–20% of the cells develop competence (Hamoen, et
al., 2003). Distinct modes of gene expression enable isogenic population of B. subtilis to diversify the
response to environmental changes (Leisner, et al., 2008).
In the case of genetic competence we spoke about competence regime. It is a time window,
in which competence can be developed under specific conditions by a single cell in the population.
It may result from the presence of specific compounds or it can be restricted to certain short periods
of time. The only exception is Neisseia gonorrhoeae, which appears to be competent all the time
Protein phosphorylation in bacteria. 7. Genetic competence – to be competent or not?
by Michał Obuchowski
33
(Sparling, 1966). When a particular cell becomes competent, over a hundred of genes become
expressed. Genes connected directly with uptake of DNA from the environment are grouped in four
com operons: comC, comE, comF, and comG. In addition to com genes, recA, nusA and addAB genes
are upregulated. Proteins encoded by comE operon form a membrane localized receptor for DNA.
Those encoded by comG form competence pseudopilus, and proteins encoded by comF – a helicaselike protein, which acts as the motor for DNA transport and is required for DNA unwinding.
The overall process of DNA transport from environment into the cell is dependent on proton motive
force of the cell (Maier, et al., 2004). All products of comG operon are synthesized as pre-proteins
and need to be processed by ComC peptidase before assembly (Kovacs, et al., 2009). Genetic
material is taken up as single-stranded DNA. Non-transforming strand is degraded extracellularly.
Incoming DNA strand is protected from nuclease by ssDNA-binding proteins, such as Ssb, DprA, or
RecA. RecA, together with AddAB proteins, forces its integration by homologous recombination.
If the transforming DNA is a plasmid, the second strand is synthesized de novo and the plasmid is
stably maintained. At an intrinsically defined switching period individual cells switch to the state of
high level of ComK, and in consequence to the state of competence. After a certain period of time,
the level of ComK decreases, and the cell loses its ability to uptake DNA. This is called the escape
from competence.
Competence can be fruitful for the cell, but it can also be dangerous. When a cell becomes
competent, DNA replication and cell division are blocked (Haijema, et al., 2001). Non-dividing cell can
be easily over-grown by competitors, so competence state should be relatively short for B. subtilis.
Gram-positive bacteria possess a thick cell wall. Penetration of DNA through such a wall is
impossible. In order to successfully uptake DNA, the wall thickness needs to be reduced. This process
occurs in response to the presence of ComEA receptor protein in the cell membrane, leading to
formation of a certain kind of a hollowing in the cell wall, by which DNA can reach the receptor.
When a cell escapes from the competence state, the thickness of the wall is restored.
Genetic competence is stimulated in the environment by co-presence of several factors, such
as nutrient limitation, growth phase, cell density and other types of stress. The regime for particular
bacterial species may greatly vary from the other, so often it is difficult to reproduce such conditions
in a laboratory. It is possible that many other bacteria may be competent in certain conditions, but
we simply cannot find them. As was said, competence is one of the sets of responses to unfavourable
conditions, but may also serve as a way of exchanging genetic material between bacteria.
34
1TProtein phosphorylation in bacteria. 7. Genetic competence – to be competent or not?
by Michał Obuchowski
8. PTS – a matter of taste
The ability of particular bacterial species to survive in the environment can be simplified to
the ability to transport and use available carbon and energy sources more efficiently than opponents.
It should be pointed out that free-living bacteria face starving conditions nearly constantly,
interrupted by rare occasions when carbon sources suitable for bio-assimilation are in excess.
In order to face these unpredictable conditions, bacteria possesses several systems of carbohydrate
uptake and utilize them in the most efficient way. Bacteria must often choose the best carbon source
from a mixture, which is actually present in the environment. To deal with this, bacteria developed
a sophisticated mechanism, which controls and coordinates carbon and energy metabolism. From
the historical point of view, the effect of presence of different sugars was described over hundred
years ago in yeast model (Dienert, 1900). Several years later, in the 1940s, the phenomenon of
diauxic growth was observed by Monod (Monod, 1942). Currently, after almost a hundred years of
research, we have drawn the picture of carbon uptake and metabolism in Gram-positive and Gramnegative bacteria. We have acquired a lot of information even at the molecular level, but still many
questions remain unanswered. Up to date, 22 systems of carbohydrate utilisation have been
identified in Bacillus subtilis (Stulke, et al., 2000). A variety of sugar uptake systems is connected with
B. subtilis ecological niche. As a soil bacterium, it has access to large amounts of polysaccharides
produced by plants. Most common of these are celluloses and chemiceluloses. These polymers
cannot be uptaken by the cells — they need to be decomposed first. In order to do this, bacteria
secrete numerous enzymes, which degrade complex nutrients to absorbable forms. It is worth of
noticing that bacteria of Bacillus genus lack enzymes necessary for degradation of cellulose, but they
can modify it to cellobiose (Priest, 1977).
Bacteria have developed a unique system of sugar transport – phosphoenolpyruvatecarbohydrate phosphotransferase system, in short called PTS. This system is specific only for bacteria
and was not found so far in Archea and Eucaryota (Warner, et al., 2003). It is important to remember
that Gram-positive and Gram-negative bacteria differ in certain aspects of PTS functioning. It will be
clear when we take a closer look at how it is organized in those organisms. Let us start with Grampositive bacteria. The core of PTS consists of EI, Hpr and enzymatic complex of EII. EI is
a well-conserved protein, present in both Gram-positive and -negative bacteria, of molecular mass
equal to approximately 63kDa. It is encoded by ptsI gene. This enzyme is capable of
autophosphorylation in the presence of Mg2+ on a conserved histidine residue, located at N-terminus
of the protein. C-terminus of EI contains a phosphoenolpyruvate (PEP) binding site and is necessary
for dimerisation (Chauvin, et al., 1996). The second PTS component, Hpr, is encoded by ptsH gene,
has molecular mass of approximately 10kDa and has been found in various organisms (Postma, et al.,
1993). This protein can be phosphorylated at His15 by EI or Ser46 by Hpr kinase/phosphatase
protein. Histidine phosphorylation site is directly linked to PTS functioning. In contrast, Ser46 residue
exhibits a regulatory function. The last component, EII complex, is a membrane-linked permease of
a particular sugar. In principle, the whole EII complex consists of four subunits: EIIA, usually
a cytoplasmic protein, which may be phosphorylated by phosphohistidine-Hpr (P-His-Hpr); EIIB,
a membrane protein facing interior of the cell, which receives the phosphate group from EIIA and
transfers it onto the carbohydrate molecule after its transport to the cytoplasm; EIIC and EIID
(if present), which are trans-membrane proteins required for translocation of a particular sugar
(Deutscher, et al., 2006). Usually, bacteria posses several EII complexes for various carbohydrates,
Protein phosphorylation in bacteria. 8. PTS – a matter of taste
by Michał Obuchowski
35
e.g. B. subtilis possesses 14 EII complexes for glucose, sucrose, β-glucosides, maltose, glucosamine,
N-acetyl-glucosamine, tethalose, oligo-β-glucoside, oligo-β-mannoside, fructose, mannitol, mannose
and cellobiose transport (Stulke, et al., 2000). Internalized sugar is converted into glucose
and metabolized by glycolytic pathway. One of the intermediates in the glycolitic pathway is fructose
1,6-biphosphate (FBP). Let us keep this molecule in mind and see how PTS works. The phosphate
group is taken from phosphoenolpyruvate (PEP) and transferred onto His15 of Hpr. Next, P-His-Hpr
passes the phosphate group to EIIA, which passes it to EIIB, and finally it is passed onto the
carbohydrate molecule. But how can the cell choose, which available carbohydrate should be taken
first? For each bacteria it is possible to prepare a hierarchal list of sugars. Usually, glucose is at the
very top. When glucose is present, phosphate coming from PEP is transferred onto internalized
molecule of sugar via PTS. This results in a relatively low level of P-His-Hpr. When P-His-Hpr is at a
low level, the amount of phosphate is insufficient for its transfer to EII permeases, other than EIIGly. In
such a situation any other sugar is not transported into the cell and cannot interact with the
appropriate repressor of the operon, which encodes genes involved in its metabolism. The simplest
example is the lac operon. If lactose cannot enter into the cell, LacI will repress lac transcription. This
mode of action is called inducer exclusion. Another way of control is induction prevention. Several
catabolic operons of B. subtilis are regulated by anti-termination. Activity of specialized
anti-terminator proteins is necessary for obtaining successful transcription. Some of these proteins
require the presence of P-His-Hpr for activity (Arnaud, et al., 1992). When Hpr phosphorylated on
histidine residue is present, the phosphate group is transferred onto SacT or LicT. A precise molecular
mechanism of regulation of SacT or LicT by phosphorylation is not known, because it has been show
in vitro that these proteins are active in non-phosphorylated form, as well as in phosphorylated
(Arnaud, et al., 1996). In addition to regulatory mechanisms described earlier, another one is
present. It is the CcpA (catabolite control protein) protein, which is a major effector of catabolite
repression in B. subtilis. CcpA is a transcription regulator, which binds to cre sequences (cataboliteresponsive sequences). Effect of CcpA binding may be different, due to localisation of cre sequences.
If cre sequences are placed inside the transcription promoter or immediately downstream of it, then
CcpA acts as a repressor. But if cre sequences are located upstream of the transcription promoter,
CcpA acts as an activator. Together, all mechanisms of controlling expression or functioning of
enzymes involved in carbohydrate uptake are called catabolite repression (CCR).
How is the action of CcpA controlled by PTS? In a pretty simple way. DNA-binding form of
CcpA is a hetero-tetramer composed of dimers of CcpA and P-Ser-Hpr protein. In addition, a paralog
of Hpr–Crh protein is present in the cell. Crh lost conserved histidine residue necessary for phosphate
transfer by PTS, but it contains a conserved serine residue and it is phosphorylated by HprK. When
Crh is phosphorylated, it may form a complex with CcpA and may act as a transcription regulator
(Stulke, et al., 2000). The HprK kinase is responsible for phosphorylation of Hpr or Crh on the serine
residue. Activity of this enzyme is directly linked with carbon status of the cell. Growth of the cells in
glucose-rich medium lead to a high level of FBP and glycerate-2-phosphate. These molecules activate
kinase activity of HprK. On the other hand, the presence of inorganic phosphate stimulates
phosphatase activity of Hpr kinase (Kravanja, et al., 1999). Activated HprK protein phosphorylates
Hpr at serine 46 of Crh. P-Ser-Hpr protein has greatly decreased affinity for EI enzyme (Deutscher, et
al., 1984). In consequence, efficiency of PTS is decreased. When the level of FBP is lowered, HprK
dephosphorylates P-Ser-Hpr and overall activity of PTS is elevated again.
36
1TProtein phosphorylation in bacteria. 8. PTS – a matter of taste
by Michał Obuchowski
When the amount of glucose available to cells is limited, the relative level of P-His-Hpr
increases, because phosphate is not transmitted to EI or further. Increased concentration of
histidine-phosphorylated Hpr can transmit phosphate onto other carbohydrate permeases, such as
EIIAGlc or glycerol kinase GlpK. Phosphorylation of other permeases than the glucose permease allows
for entry of these sugars into the cell and for the release of repression of its catabolite operons.
At the same time, Hpr kinase stimulated by increased concentration of inorganic phosphate,
dephosphorylates Hpr protein and ends catabolic repression caused by CcpA and Crh proteins.
Due to the fact that CcpA exhibits different affinity to various cre elements, activation of catabolite
operons is ordered from the most preferred carbohydrate to the less preferred. It should be pointed
out, that loss of CcpA or cre site only results in reduced catabolite repression, because multiple
mechanisms of CCR exist in the cell (Dahl, et al., 1995).
In Gram-negative bacteria the overall scheme of PTS action is identical. The major difference
is that bacteria such as E. coli have no Hpr kinase. They obviously possess Hpr protein, but it is
phosphorylated only on histidine residue. The regulatory function is taken by EIIAGlc protein. When
cells have enough glucose available, EIIAGlc is mostly in the dephosphorylated form. In this state it can
bind to other carbohydrate permeases and inhibit their activity. In the same way EIIAGlc is bound to
glycerol kinase, which results in its inhibition. When glucose becomes unavailable, the intracellular
concentration of phosphorylated EIIAGlc increases. P-EIIAGlc forms a complex with adenylate cyclase
and stimulates its activity. It has been postulated that phosphorylated EIIAGlc does not directly
interact with adenylate cyclase and that an additional protein is required to stimulate its activity.
In consequence, intracellular concentration of cAMP increases. In Gram-negative bacteria Crp
protein is present instead of CcpA protein. You have probably heard about this protein for the first
time, when you learnt about the regulation of the lac operon. Crp is one of the truly global
regulators, which control expression of a large number of genes and operons. Crp is activated by
binding of cAMP. As was the case with CcpA in Gram-positive bacteria, different genes or operons
require different level of the Crp-cAMP complex (Deutscher, et al., 2006).
PTS system is also connected with other cellular responses. For example, it has been noticed
that E. coli shows chemotaxis towards many PTS sugars. Upon closer examination it was established
that this chemotactic response is precisely connected with transport of carbohydrates, but not with
their binding or metabolism (Grisafi, et al., 1989). Another connection between PTS and chemotaxis
is the fact that unphosphorylated EI enzyme can interact with CheA kinase and that it decreases its
activity (Lux, et al., 1995). Lowered activity of CheA results in longer periods of smooth swimming of
the cell. In B. subtilis the situation is different. In this bacterium, PTS and chemotaxis are linked by
the methyl-accepting chemotaxis protein McpC (Muller, et al., 1997). In B. subtilis also the activity of
CheA kinase is regulated by EI, but in this case the phosphorylated form is active in terms of CheA
binding. It looks strange, but you should remember that chemotactic response regulator, CheY, has
an opposite effect on swimming behaviour in E. coli and B. subtilis. Phosphorylated CheY leads to
tumbling in E. coli and to smooth swimming in B. subtilis. Having that in mind, it is clear that response
of both bacteria to the presence of PTS sugars is the same – smooth swimming (Deutscher, et al.,
2006).
Bacteria have evolved sophisticated systems of sensing the nutritional situation and responding accordingly. These systems allow the cell to maximize growth rate and possibly to overgrow the
competitors. Carbon status detected by PTS activity is connected with several bacterial
developmental programs, such as chemotaxis, sporulation and virulence. Several examples
Protein phosphorylation in bacteria. 8. PTS – a matter of taste
by Michał Obuchowski
37
describing direct link between PTS and expression of virulence genes can be found in the literature
(Deutscher, et al., 2005). PTS and its connections with other metabolic pathways is another example
of complex regulation of gene expression and enzymatic activity, which takes place in bacterial cells.
38
1TProtein phosphorylation in bacteria. 8. PTS – a matter of taste
by Michał Obuchowski
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