Download Vipp1: a very important protein in plastids?!

Journal of Experimental Botany, Vol. 63, No. 4, pp. 1699–1712, 2012
doi:10.1093/jxb/err357 Advance Access publication 29 November, 2011
REVIEW PAPER
Vipp1: a very important protein in plastids?!
Ute C. Vothknecht1,*, Stephanie Otters1, Raoul Hennig2 and Dirk Schneider2,*
1
2
Department of Biology I, LMU Munich, D-82152 Planegg-Martinsried, Germany
Department of Pharmacy and Biochemistry, Johannes Gutenberg-University Mainz, D-55128 Mainz, Germany
* To whom correspondence should be addressed. E-mail: [email protected] or [email protected]
Received 5 September 2011; Revised 13 October 2011; Accepted 14 October 2011
Abstract
As a key feature in oxygenic photosynthesis, thylakoid membranes play an essential role in the physiology of plants,
algae, and cyanobacteria. Despite their importance in the process of oxygenic photosynthesis, their biogenesis has
remained a mystery to the present day. A decade ago, vesicle-inducing protein in plastids 1 (Vipp1) was described to
be involved in thylakoid membrane formation in chloroplasts and cyanobacteria. Most follow-up studies clearly
linked Vipp1 to membranes and Vipp1 interactions as well as the defects observed after Vipp1 depletion in
chloroplasts and cyanobacteria indicate that Vipp1 directly binds to membranes, locally stabilizes bilayer structures,
and thereby retains membrane integrity. Here current knowledge about the structure and function of Vipp1 is
summarized with a special focus on its relationship to the bacterial phage shock protein A (PspA), as both proteins
share a common origin and appear to have retained many similarities in structure and function.
Key words: Chloroplasts, cyanobacteria, PspA, thylakoid membrane biogenesis, vesicle transport, Vipp1.
Emergence of Vipp1 as a factor for thylakoid
biogenesis
Thylakoids are characteristic structures of cyanobacteria
and chloroplasts, and the major complexes of oxygenic
photosynthesis are embedded in these membranes. The
thylakoid membrane system evolved in cyanobacteria in
close correlation with the evolution of oxygenic photosynthesis and generated a novel bacterial compartment, the
thylakoid lumen. Thylakoids are exclusively linked to
organisms carrying out oxygenic photosynthesis, and the
facility to build up and to modify such a complex membrane system has been transferred from the cyanobacterial
ancestor to the plant cell during the endosymbiotic establishment of chloroplasts (reviewed in Vothknecht and
Westhoff, 2001). Thylakoid membrane biogenesis is a multidimensional process, involving a concerted interplay of lipid,
protein, and pigment synthesis, as well as their transport and
integration into functional membrane-integral complexes. It
is still unclear whether they assemble completely de novo or
whether their membrane scaffold derives from the inner
envelope of chloroplasts or the plasma membrane of
cyanobacteria, respectively. While insertion of photosynthetic proteins into already existing thylakoid lipid bilayers
has been analysed in some detail (reviewed in Aldridge
et al., 2009), there is very little information about components involved in the formation of the thylakoid membrane
itself (reviewed in Vothknecht and Westhoff, 2001; Nickelsen et al., 2010).
The vesicle-inducing protein in plastids 1 (Vipp1) was first
described in 1994 as a protein located in the chloroplasts of
Pisum sativum (Li et al., 1994). Vipp1 is nuclear encoded
and contains an N-terminal sequence sharing common
characteristics with other chloroplast-directed transit peptides. Thus, like most plastidal proteins, Vipp1 is translated
as a precursor on cytosolic ribosomes and is afterwards
imported into the chloroplast by the TOC–TIC general
import pathway (Schnell et al., 1997). An initial analysis
further suggested that the protein is located at the inner
envelope membrane as well as the thylakoids. Due to its
localization and its mature size of 30 kDa, the protein was
Abbreviations: PG, phosphatidyl glycerol; PS, photosystem; Psp, phage shock protein; Sec, general secretory pathway; Tat, twin-arginine translocation pathway; TIC,
translocon on the inner chloroplast membrane; TOC, translocon on the outer chloroplast membrane; Vipp1, vesicle-inducing protein in plastids 1.
ª The Author [2011]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
For Permissions, please e-mail: [email protected]
1700 | Vothknecht et al.
first named M30 (membrane-associated protein of 30 kDa)
(Li et al., 1994). However, Vipp1 does not display any
features of intrinsic membrane proteins, as was revealed
by structural analysis and supported by experimental
approaches (Li et al., 1994; Kroll et al., 2001). Instead,
further studies demonstrated a rather peripheral association
of Vipp1 with chloroplast membranes. The unique localization of Vipp1 at the two membranes bordering the stromal
compartment led to the presumption that Vipp1 might
somehow be involved in the assembly of the thylakoid
membrane system (Li et al., 1994). More precisely, it has
been suggested that the protein is involved in the transport
of lipids, which are synthesized at the chloroplast inner
envelope. A potential involvement of Vipp1 in thylakoid
membrane biogenesis was supported several years later by
the drastic phenotype of an Arabidopsis thaliana high
chlorophyll fluorescence mutant line (hcf155), in which the
expression of the vipp1 gene was strongly reduced due to
a T-DNA insertion in its promoter region (Kroll et al.,
2001). Mutant plants expressed only about a fifth of the
Vipp1 wild-type protein content under normal growth
conditions and were incapable of autotrophic growth. Even
on sugar-supplemented growth medium they showed
a strong albinotic phenotype and did not survive beyond
the stage of a small rosette (Kroll et al., 2001). Transmission
electron microscopy of leaf sections revealed that mutant
plants were not able to build a properly structured
thylakoid membrane system. Their chloroplasts, while
almost normal in size, contained very few internal membranes in rather severely distorted structures that are
diffusely distributed inside the organelle. Correlating with
their lack of thylakoid membranes, the capacity for
oxygenic photosynthesis was extremely decreased (Kroll
et al., 2001). Furthermore, hcf155 mutant plants displayed
another phenotype in accordance with Vipp1’s alleged
function in lipid transfer. Formation of vesicular structures
at the inner envelope of chloroplasts could no longer be
observed in the mutants, indicating that Vipp1 might be
essential for this feature. Thus, the protein was renamed
‘vesicle-inducing protein in plastids 1’ (Kroll et al., 2001).
At the same time, a vipp1 mutant of Synechocystis sp. PCC
6803 was described. While not fully segregated, mutant cells
had a drastically reduced Vipp1 content. This cyanobacterial mutant displayed a similar phenotype to the hcf155
mutant line, showing a comparable loss of thylakoid
membrane content and structure, as well as reduced photosynthetic activity (Westphal et al., 2001a). These findings not
only supported the function of Vipp1 independently in two
different organisms but also indicated a phylogenetically
conserved function.
PspA and Vipp1, a shared past (and
present?)
Phylogenetic analyses strongly support Vipp1 as a protein
of prokaryotic origin that is found exclusively in cyanobacteria and chloroplasts (Li et al., 1994; Kroll et al., 2001;
Westphal et al., 2001a; Fuhrmann et al., 2009a). This
indicates a development in parallel to the emergence of the
thylakoid membrane system and a connection to the
evolution of oxygenic photosynthesis. However, proteins
with significant similarities to Vipp1 have been identified in
several non-photosynthetic bacteria (Fig. 1) in form of the
phage shock protein A (PspA) (Brissette et al., 1990). PspA
was first characterized following infection of Escherichia coli
cells with filamentous phages, whereby the phage gene pIV
protein was found to induce pspA expression to extremely
high levels (Brissette et al., 1990). It was subsequently
shown that PspA is a member of a bacterial stress response
system, referred to as the ‘phage shock protein system’ (Psp
system) (Brissette et al., 1991). The Psp system in E. coli
responds not only to the invasion of filamentous phages but
also to a number of different extracytoplasmic stress
factors, such as heat, hyperosmotic conditions, organic
solvents (Brissette et al., 1991; Weiner and Model, 1994;
Kobayashi et al., 1998), inhibition of lipid biosynthesis
(Bergler et al., 1994), and mislocalization or blockage of the
protein export machinery (Kleerebezem and Tommassen,
1993; Kleerebezem et al., 1996; DeLisa et al., 2004). A
common denominator of these stress conditions is the
reduction of the proton motive force across the cytoplasmic
membrane. In total, the Psp system in E. coli comprises
seven psp genes. In addition to the pspABCDE operon, the
transcriptional activator PspF as well as the effector protein
PspG are encoded by separate genes (Brissette et al., 1991;
Elderkin et al., 2002; Lloyd et al., 2004). Furthermore, the
E. coli two-component system consisting of ArcA and ArcB
has been shown to be involved in signal sensing and
induction of the Psp-mediated stress response (Jovanovic
et al., 2006, 2009).
PspA is a hydrophilic protein of 25 kDa that, like Vipp1,
shares no features of integral membrane proteins but
appears to be peripherally associated with the cytoplasmic
membrane of bacterial cells (Brissette et al., 1990; Cserzo
et al., 1997; Kobayashi et al., 2007). As the most abundant
protein and primary effector of the Psp response, PspA is
responsible for several phenotypes connected to this system
(Weiner and Model, 1994; Kleerebezem et al., 1996; DeLisa
et al., 2004). It is assumed that PspA supports the bacterial
cell in stress situations by blocking proton leakage
(Kobayashi et al., 2007) or by inducing the down-regulation
of proton motive force-consuming processes (Engl et al.,
2009). The Psp system appears to be widespread throughout
all bacterial domains. Psp proteins have been identified in
Gram-negative and Gram-positive bacteria, as well as in
archaea and cyanobacteria. In Yersinia enterocolitica, where
the stress response is well established, the Psp system was
found to be induced by two outer membrane secretins.
Furthermore, it plays an important role in maintaining the
membrane stability of the pathogen during cell infection
(Darwin and Miller, 2001). Psp proteins are also highly
expressed in Salmonella enterica (Eriksson et al., 2003) and
Shigella flexneri (Lucchini et al., 2005) during macrophage
infection. A functional analysis of the Psp response in the
halophilite archaeon Halifax volcanii has pointed out that
Vipp1: a very import protein in plastids | 1701
Fig. 1. Phylogenetic tree construction of PspA and Vipp1. Included in the analysis were proteins with sequenced similarity to PspA and
Vipp1 from Actinomyces odontolyticus (Aco), Alteromonas macleodii (Alm), Anabaena variabilis (Anv1, Anv2), Arabidopsis thaliana (Ath),
Arthrobacter chlorophenolicus (Arc), Azospirillum sp. (Azs), Bacillus sp. (Bas), Bacillus subtilis (Bacs), Bifidobacterium animalis (Bia),
Bifidobacterium gallicum (Big), Burkholderia phymatum (Bup), Caldicellulosiruptor bescii (Cab), Caldicellulosiruptor owensensis (Cao),
Candidatus Korarchaeum cryptofilum (Cac), Chlamydomonas reinhardtii (Chr), Chlorella variabilis (Chv), Chloroherpeton thalassium (Cht),
Citrobacter sp. (Cis), Clostridium botulinum (Clb), Clostridium sp. (Cls), Crocosphaera watsonii (Crw1, Crw2), Cyanothece sp. (Cys1,
Cys2), Delftia sp. (Des), Desulfatibacillum alkenivorans (Desa), Desulfobacterium autotrophicum (Dea), Desulvovibrio vulgaris RCH1 (Dev),
Desulvovibrio vulgaris DP4 (Desv), Ectocarpus siliculosus (Ecs), Eikenella corrodens (Eic), Enterobacter sp. (Ens), Erythrobacter sp. (Ers),
Escherichia coli (Esc), Gloeobacter violaceus (Glv), Gluconobacter oxydans (Glo), Glycine max (Glm), Haliangium ochraceum (Hao),
Haloarcula marismortui (Ham), Haloquadratum walsbyi (Haw), Halorhabdus utahensis (Hau), Haloterrigena turkmenica (Hat), Heliobacterium modesticaldum (Hem), Hirschia baltica (Hib), Hordeum vulgare (Hov), Hypomonas neptunicum (Hyn), Klebsiella pneumoniae
(Klp), Kytococcus sedentarius (Kys), Lyngbya majuscule (Lym1, Lym2), Lyngbya sp. (Lys1, Lys2), Micrococcus luteus (Mil), Microcoleus
vaginatus (Miv1, Miv2) Micromonas pusilla (Mip), Micromonas spec. (Mis), Myxococcus xanthus (Myx), Natrialba magadii (Nam), Neisseria
subflava (Nes), Nodularia spumigena (Nods1, Nods2), Nostoc punctiforme (Nop1, Nop2), Nostoc sp. (Nos1, Nos2) Novosphingobium
sp. (Novs) Oryza sativa (Ors), Oscillatoria sp. (Oss1, Oss2), Phaeodactylum tricornutum (Pht), Photobacterium profundum (Phop),
Phyllostachys edulis (Phe), Physcomitrella patens (Php), Picea sitchensis (Pics), Pisum sativum (Pis), Planctomyces brasiliensis (Plb),
Planctomyces maris (Plm), Plesiocystis pacifica SIR-1 (Plp, Plep) Populus trichocarpa (Pot), Prochlorococcus marinus (Prm), Proteus
mirabilis (Prs), Pseudomonas syringae (Pss), Renibacterium salmoninarum (Res), Rhodospirillum rubrum (Rhr), Ricinus communis (Ric),
Ruminococcus sp. (Rus), Salmonella enterica (Sae), Selaginella moellendorffii (Smo), Shewanella baltica (Shb), Sorangium cellulosum
(Soc), Sorghum bicolour (Sob), Sphingobium japonicum (Spj), Streptomyces lividans (Stl), Streptomyces pristinaespiralis (Stp),
1702 | Vothknecht et al.
pspA expression was up-regulated in response to salt stress
(Bidle et al., 2008). Phylogenetic analysis indicates that
a minimal functional Psp system—containing just PspA, B,
C, and F—is conserved in a-, d- and c-proteobacteria,
whereas other Psp components show much less conservation or are even absent (Darwin, 2005). Moreover, PspA
homologues also appear in organisms without connection
to any other component of the Psp response, suggesting
either a strong reduction of the Psp system or a newly
acquired function of the remaining PspA homologue
(reviewed in Darwin, 2005). In Streptomyces lividans,
expression of pspa is strongly induced under membrane
stress (Vrancken et al., 2008), which indicates that PspA has
retained an important effector function in the stress response of this Gram-positive bacterium despite the loss of
other Psp members. LiaH, the PspA homologue of the
Gram-positive bacterium Bacillus subtilis (Mascher et al.,
2004), belongs to the Lia system that is conserved in all
Firmicutes, Bacillus, and Listeria species, where it regulates
the bacterial stress response. The regulatory Lia system is
comprised of the LiaRS two-component system linked to
LiaF, a membrane-anchored negative regulator of LiaRdependent gene expression. The LiaRS two-component
system recognizes different antibiotics and triggers a 100to 1000-fold induction of the liaIHGFSR operon. Beside
a strong induction in response to external cell wall antibiotics that affect the lipid II cycle (Mascher et al., 2004;
Pietiainen et al., 2005), expression of the Lia system is also
induced by alkaline shock, as well as by treatment with
organic solvents and secretion stress (Petersohn et al., 2001;
Mascher et al., 2004; Hyyrylainen et al., 2005). The Lia
system of B. subtilis thus seems to represent a complex
regulatory stress response system in Gram-positive bacteria
comparable with the Psp response system found in E. coli
(Wolf et al., 2010). While most components of the Psp and
Lia system differ, the primary target proteins of both
systems, LiaH and PspA, show sequence and structural
homology, indicating a similar mode of action.
Many cyanobacterial genomes possess two genes encoding proteins with significant homology to members of the
PspA/Vipp1 protein family (Fig. 1), even if they lack other
genes encoding proteins of the original Psp response. Based
on a phylogenetic analysis, these proteins divide into two
different clades, suggesting that cyanobacteria require PspA
as well as Vipp1. In the genome of Anabaena sp. PCC 7120,
the genes encoding these two proteins are localized adjacent
to each other with a gap of only 162 bp between the stop
codon of the former and the start codon of the latter
(Kaneko et al., 2001). A similar arrangement can be found
in other cyanobacteria, suggesting that Vipp1 has evolved in
an ancestor of today’s cyanobacteria from PspA by gene
duplication and was passed to the plant lineage by the
cyanobacterial endosymbiont (Westphal et al., 2001a). No
pspA gene has been identified explicitly in any plant or alga,
indicating that PspA was either absent in the original
endosymbiont or was lost in the plant lineage shortly after
the endosymbiotic event. While cyanobacterial PspA proteins cluster together with the Vipp1 branch containing both
cyanobacterial and plant Vipp1 proteins, PspA proteins
from bacteria other than cyanobacteria cluster in a separate
clade, which also includes PspA proteins from Archaea
(Fig. 1). This strongly supports the hypothesis that Vipp1
evolved from a cyanobacterial PspA rather than originating
in a different bacterial ancestor.
Initially, the situation appeared rather straightforward,
with non-photosynthetic bacteria possessing only PspA,
chloroplasts only Vipp1, and cyanobacteria encoding both
proteins (Fig. 1). This observation linked the presence of
Vipp1 to thylakoid membranes and supported a thylakoidspecific function. Such an assumption was further supported by the apparent lack of a vipp1 gene in the genome
of Gloeobacter violoceus sp. PCC 7421 (Nakamura et al.,
2003), a unicellular, obligate photoautotrophic bacterium
that branches at an early stage of cyanobacterial evolution
(Nelissen et al., 1995; Honda et al., 1999). It contains
structural features that differ from those of common
cyanobacteria, such as the lack of thylakoid membranes,
and its photosynthetic machinery is located on the cytoplasmic membrane (Rippka et al., 1974). As Gloeobacter
provides several primordial characteristics, the organism is
an important tool to investigate the origin of oxygenic
photosynthesis (Nakamura et al., 2003; Mimuro et al.,
2008). In agreement with the alleged function of Vipp1 in
thylakoid biogenesis, its absence in Gloeobacter provided
a simple explanation for the inability of this organism to
build up a functional thylakoid membrane system. Gloeobacter does contain a protein of the Vipp1/PspA family,
which is annotated as a PspA homologue due to certain
conserved structural characteristics (see below). However, in
phylogenetic analysis, this Gloeobacter protein is placed on
a separate side branch of the cyanobacterial Vipp1/PspA
clade, so that a definite phylogenetic relationship to either
PspA or Vipp1 cannot be obtained (Fig. 1). The availability
of many more completely sequenced cyanobacterial
genomes has caused the situation to become more complex.
The genomes of several cyanobacteria, such as some
Prochlorococcus species, also lack a gene encoding Vipp1,
despite the presence of thylakoids in these organisms
(Dufresne et al., 2003). In some of these cyanobacteria, the
supposed lack of Vipp1 might be due to a misannotation of
Synechococcus sp. (Syns1, Syns2), Synechocystis sp. (Sys1, Sys2), Thalassiosira pseudonana (Thp), Thiomonas intermedia (Thi),
Trichodesmium erythraeum (Tre1a, Tre1b, Tre2), Vibrio sp. (Vibs), Vitis viniferum (Viv), Volvox carteri (Voc), Yersinia pestis (Yep), and
Yersinia enterocolitica (Yee). Sequence alignments were obtained by Clustal 1.8 (Thompson et al., 1997). Phylogenetic tree construction
was performed by maximum likelihood using Phyml 3.0 (Guindon and Gascuel, 2003). Transit sequences were omitted from the plant
sequences for the analysis.
Vipp1: a very import protein in plastids | 1703
the vipp1 genes as pspA, but in some of them Vipp1 indeed
appears to be absent. While these findings question the
direct correlation of Vipp1 to thylakoid membrane evolution, it is possible that these organisms have secondarily lost
Vipp1 and now utilize a different system for thylakoid
membrane biogenesis and/or maintenance.
Since Vipp1 originated from PspA, the question arose as
to what extent PspA function is still conserved in Vipp1
proteins. Expression of Vipp1 from the cyanobacterium
Synechocystis sp. PCC6803 in an E. coli DpspA deletion
strain prevented blockage of protein translocation (DeLisa
et al., 2004), indicating that the cyanobacterial Vipp1 could
somehow functionally replace the bacterial PspA. In
contrast, endogenous PspA present in a Synechocystis
Dvipp1 depletion strain was not sufficient to compensate
the deficiency causing the drastic phenotype of those
mutants, and thus to take over the function of Vipp1
(Westphal et al., 2001a; Fuhrmann et al., 2009b; Gao and
Xu, 2009). Obviously, PspA is not necessary for the
bacterial cells in the absence of stress factors, since E. coli
DpspA mutants are perfectly viable (Brissette et al., 1991).
In contrast, vipp1 mutants from Arabidopsis and Synechocystis show a dramatic phenotype even if vipp1 expression is
not completely abolished but merely strongly decreased
(Kroll et al., 2001; Westphal et al., 2001a; Aseeva et al.,
2007; Gao and Xu, 2009). Along with the fact that many
cyanobacteria still possess both proteins, there is strong
evidence that Vipp1 proteins have gained a novel function
in cyanobacteria and chloroplasts. However, Vipp1 has also
been clearly correlated to stress response. In Synechocystis,
Vipp1 was found amongst those plasma membrane proteins
exhibiting the highest enhancements under high salt conditions (Huang et al., 2006). In contrast, the vipp1 gene was
significantly down-regulated in the cold (Suzuki et al.,
2001). Also, the Chlamydomonas Vipp1 was found in
a group of proteins whose expression was up-regulated
under high light (Im and Grossman, 2002). Nevertheless, in
all these cases it is not clear whether the change in vipp1
expression represents a specific stress response or whether it
is due to a general function of Vipp1 in thylakoid
membrane synthesis and maintenance, which is affected
under these conditions.
PspA and Vipp1 proteins display certain conservation in
primary amino acid sequence but, even in the same
organism, such as in Synechocystis sp. PCC 6803, they share
just ;30% sequence identity and 50% sequence similarity
(Bultema et al., 2010). Nevertheless, all PspA/Vipp1 family
members exhibit a common N-terminal core structure, the
so-called PspA domain that is ;220 amino acids in length
(Westphal et al., 2001a). Furthermore, the secondary and
tertiary structures as well as the intrinsic propensity of PspA
and Vipp1 to form higher ordered oligomers appear to be
highly conserved (Aseeva et al., 2004; Hankamer et al., 2004;
Fuhrmann et al., 2009a; Bultema et al., 2010).
Based on computational predictions and spectroscopic
measurements, an essentially exclusively a-helical structure
was identified for the PspA-like domain of Vipp1/PspA
proteins (Fig. 2A) (Hankamer et al., 2004; Fuhrmann et al.,
2009a; Bultema et al., 2010). Only few amino acids within
this region are predicted to be unstructured and to interrupt
the helical domain. In contrast to PspA, Vipp1 proteins are
longer due to a random coil spacer of variable size,
connecting the PspA-like domain to a short a-helical
domain of ;30 amino acids (Westphal et al., 2001a). This
defined a-helical extension at the C-terminus appears to be
a characteristic feature of Vipp1 proteins that clearly
distinguishes Vipp1 from PspA.
The a-helical regions of the PspA domain have a high
propensity to form coiled-coil structures and to homooligomerize to higher ordered rings with molecular masses
>1 MDa. While the exact tertiary structure of Vipp1 is
presently unknown, large parts of the coiled-coil domain are
The structure of Vipp1
Vipp1 proteins of plants and algae exhibit an N-terminal
extension that has evolved as a transit peptide to ensure the
import of the cytosolic-translated protein into the chloroplast (Li et al., 1994; Westphal et al., 2001a). Since the
transit peptide is removed after translocation, it has no
bearing on the structure of the mature protein and is
therefore mostly excluded in structural comparisons between Vipp1 and PspA. While the phylogenetic relationship
between Vipp1 and PspA is evident, it remains unclear to
what extent the function of Vipp1 was altered in the course
of evolution and which changes within the protein might
affect any difference in function between the two proteins.
Fig. 2. Vipp1 oligomer formation. (A) Schematic of the Synechocystis Vipp1 protein sequence and the positions of the helical
domains (HDs) according to regions defined for E. coli PspA
(Elderkin et al., 2005). Putative functions of the individual domains
in oligomerization are indicated. Amino acid residues 222–267
represent the C-terminal extension typical for Vipp1 proteins.
Predicted a-helical regions are indicated by grey bars.
(B) Formation of Vipp1 rings from Vipp1 monomers involves
intermediate dimeric, tetrameric, and string-like states.
(C) Individual Vipp1 rings eventually interact to form double rings
and rod-like structures.
1704 | Vothknecht et al.
involved in oligomerization (Aseeva et al., 2004; Bultema
et al., 2010). Based on structural analysis, PspA of E. coli
was subdivided into four helical domains HD1–HD4 and, in
a fragmentation approach, oligomerization of truncated E.
coli PspA proteins was analysed in detail (Elderkin et al.,
2005; Joly et al., 2009) (Fig. 2A). The centre regions HD2
and HD3 alone support formation of a PspA hexamer,
whereas the HD1–HD2–HD3 protein forms dimers. Thus,
while HD2 and HD3 are critical for hexamer formation,
HD3 negatively influences hexamerization, thereby however
stabilizing a dimeric PspA structure. HD4 supports formation of a PspA 36mer ring structure, and, thus, this domain
appears to connect individual dimers. It might be assumed
that similar regions control assembly and disassembly of
Vipp1 oligomers in vivo, as indicated in Fig. 2A, but this has
not been assessed in detail yet. Accessibility of the individual
helical regions and their involvement in defined intermonomer contacts might be the key for controlling the actual
oligomeric state of PspA/Vipp1 proteins. Chaperones might
further be crucially involved in shielding defined regions of
the protein for controlled Vipp1 assembly/disassembly (as
discussed below).
While the tertiary structures of PspA and Vipp1 are not
well defined, the quaternary structures of both have been
analysed to a certain degree in recent years. Both proteins
form large homo-oligomeric rings, albeit with different
dimensions (Aseeva et al., 2004; Hankamer et al., 2004; Liu
et al., 2007; Fuhrmann et al., 2009a; Bultema et al., 2010).
PspA from E. coli forms a single, distinct ring structure with
a 9-fold rotational symmetry and a calculated mass of
;1 MDa. It has been suggested that this ring assembles
from nine tetrameric building blocks (Hankamer et al.,
2004). The PspA ring has an outer diameter of 20 nm, an
inner diameter of 9.5 nm, and a height of ;8.5 nm
(Hankamer et al., 2004). In contrast, Vipp1 proteins of
Arabidopsis and Synechocystis organize into variable ring
structures with molecular masses of up to ;2 MDa (Aseeva
et al., 2004; Fuhrmann et al., 2009a). While for Synechocystis Vipp1 at least six different ring structures with a 12–17
internal rotational symmetry and outer diameters between
25 nm and 33 nm have been identified, all rings have
a uniform height of 22 nm (Fuhrmann et al., 2009a). Based
on recent results it seems likely that multiple copies of
monomers (or smaller oligomeric building blocks) assemble
into string-like structures representing intermediates during
ring assembly or disassembly. Such an assumption is
supported by the observation that some string-like structures have indeed been observed in the case of the
Synechocystis protein. Furthermore, controlled disassembly
of Vipp1 results in formation of distinct tetrameric and
dimeric (dis)assembly intermediates, indicating that Vipp1
dimers are the minimal building blocks of the Vipp1 rings
(Fuhrmann et al., 2009a). At higher concentrations, individual Vipp1 rings eventually stack together and form
barrel-like double-ring structures, as observed with Vipp1
from cyanobacteria and higher plants (Aseeva et al., 2004;
Fuhrmann et al., 2009a). Such double-ring structures
potentially represent an assembly intermediate during
formation of multiple-ring stacking structures with a rodlike shape, as detected after heterologous expression and in
vitro refolding of the Chlamydomonas reinhardtii Vipp1 (Liu
et al., 2007). Such scaffold-like structures have also been
described for PspA (Standar et al., 2008). Figure 2B and C
summarize all presently identified Vipp1 structures and
a potential assembly/disassembly pathway.
The characteristic C-terminal extension of Vipp1 proteins
does not appear to be essential for the formation of rings or
rods (Aseeva et al., 2004; Liu et al., 2007). Fusing a protein
to the Vipp1 C-terminus does not interfere either with its
oligomerization propensity or with thylakoid biogenesis in
Arabidopsis (Aseeva et al., 2007). Thus, the C-terminus is
probably not crucially involved in formation of Vipp1
oligomers and a free C-terminus is not essential for Vipp1
activity. However, as the C-terminal extension appears to be
a characteristic feature of Vipp1, it is crucial for discriminating PspA from the PspA-like Vipp1 proteins.
Based on a sequence alignment of Vipp1 homologues of
cyanobacteria, algae, and higher plants, the following
consensus sequences of the Vipp1 C-terminus was generated:
V/I/L-D/E-X-E-L-E/D-E/D-L-R/(K)-R/K-X-L/I-D/
N-X-(L)-X3 (Fig. 3A). Even though the amino acids are
not strictly conserved at most positions, the properties of
individual amino acid side chains are preserved at various
places. The Vipp1 C-terminus is predicted to form an
amphiphilic a-helix with one non-polar, hydrophobic face,
containing two acidic residues at the N-terminus and two
basic residues at the C-terminus (Fig. 3B). As the
C-terminus most probably protrudes out of the ring
structures (Aseeva et al., 2004), these conserved features of
the Vipp1 C-terminal helix eventually allow and control
defined interactions with proteins, lipids, or membranes.
Structural dynamics, interactions, and
functions
Most data somehow link both PspA and Vipp1 to membranes. In chloroplasts, Vipp1 is associated with the inner
envelope as well as with thylakoid membranes, and in
cyanobacteria the protein localizes in close vicinity to both
the cytoplasmic and thylakoid membranes (Srivastava et al.,
2005; Fuhrmann et al., 2009a). Detailed studies on membrane
attachment revealed that the PspA protein of E. coli is able to
bind directly to membrane lipids, having a clear preference for
lipids with negatively charged head groups (Kobayashi et al.,
2007). In vivo, however, PspA is mainly recruited to membranes by the membrane-integral proteins PspB and C, and it
has been suggested that these proteins sense stress signals and
modulate PspF activity by a controlled interaction with PspA
(Joly et al., 2010). Only upon release of PspA from the PspBC
complex the interaction of PspA with the transcription
activator PspF permitted. This releases the suppression of the
psp operon by PspF and results in overexpression of PspA.
Subsequently, PspA binds to membranes directly for its
function in stress protection.
Vipp1: a very import protein in plastids | 1705
Fig. 3. Consensus sequence of the Vipp1 C-terminus. (A) Alignment of Vipp1 sequences from cyanobacteria, green algea, brown algae,
and plants revealed a conserved consensus sequence within the last 44 amino acids, forming the a-helical Vipp1 C-terminus. The
degree of conservation is shown by the number of bits (Crooks et al., 2004). The approach disclosed a strict conservation of side chain
properties in several positions, even if the amino acids were not conserved. (B) Helical wheel projection of the consensus sequence
underlined in A predicts the formation of an amphiphatic a-helix with a hydrophilic face on one side of the helix (indicated). Furthermore,
the helix is bookended by two conserved acidic residues at the N-terminus and two basic residues at the C-terminus. Amino acid
properties are indicated: magenta, acidic residue; blue, basic residue; black/grey, non-polar residue.
Different oligomeric structures of PspA appear to have
different functions in E. coli. When interacting with PspF,
PspA is present as a hexamer, whereas formation of the
PspA 36mer appears to be required for direct membrane
attachment and maintenance of membrane integrity. In
contrast to PspA, no hexameric Vipp1 structure has been
reported, and, as Vipp1 does not seem to interact with any
component similar to PspF, a hexameric assembly stage
might not exist. Extraction of Vipp1 from cyanobacterial
thylakoids furthermore suggests that the protein binds as
rings to membranes (Fuhrmann et al., 2009a). Soluble
Vipp1 rings have been identified in the cyanobacterial
cytoplasm, which might be less stable than membranebound rings (Fuhrmann et al., 2009a). While for PspA
membrane attachment appears to be regulated by the
oligomeric state, it is not clear how this is regulated for
Vipp1. It is noteworthy that in cyanobacteria and plants, two
Vipp1 forms showing an ;2 kDa difference in apparent
molecular mass have been identified (Kroll et al., 2001;
Aseeva et al., 2004; Liu et al., 2005; Fuhrmann et al., 2009a).
While the difference between these two Vipp1 forms has not
been determined yet, Vipp1 might be post-translationally
modified. Thus, the two Vipp1 forms observed on SDS–gels
might represent two structurally different Vipp1 populations
having different membrane binding affinities.
Individual Vipp1 monomers have an intrinsic propensity
to homo-oligomerize and to aggregate. In an Arabidopsis
mutant strain containing a massively decreased Vipp1
content, oligomers have not been observed, and thus it
seems that a critical Vipp1 concentration is needed for
complex formation (Aseeva et al., 2007). At higher concentrations, rod-like structures form, which have been suggested to represent a chloroplastic microtubule-like system
(Liu et al., 2007). Rod-like structures of unknown origin
have indeed been observed in chloroplasts as well as in
cyanobacteria (Anderson et al., 1973; Lawrence et al., 1984;
van de Meene et al., 2006), and it is therefore intriguing to
speculate that such cytoskeleton-like elements could be used
as tracks for a directed transport of proteins and/or vesicles
to distinct locations within chloroplasts or cyanobacteria.
Nevertheless, while Vipp1 rings stack at higher concentrations, no rod structures have been reported thus far for
PspA proteins. Formation of rods, which are essentially
identical to Vipp1 rods, is also observed in vitro by the ringforming, membrane-integral Holine protein of bacteriophage k (Savva et al., 2008). Since artificial formation of
1706 | Vothknecht et al.
the identified rod structures here strongly depended on the
experimental conditions, the existence of Vipp1 rods in vivo
still has to be proven unambiguously.
Oligomerization and membrane attachment of Vipp1
might be controlled by a chaperone-mediated disassembly
of Vipp1 rings. Shielding defined regions of the PspA
domain would allow controlled assembly/disassembly of
Vipp1 rings (compare Fig. 2A) and consequently membrane
binding of the protein. Vipp1 of the green alga C. reinhardtii
interacts with the chloroplast chaperone Hsp70B, as well as
with the Hsp40 homologue CDJ2 (Liu et al., 2005). Binding
of these chaperones to Vipp1 resulted in disassembly of
large Vipp1 rings and rods in vitro (Liu et al., 2007),
although it is not yet clear whether the rings disassemble
into monomers or to lower ordered oligomeric states. Thus,
the Hsp70/Hsp40 system might be part of a cellular system
which controls assembly/disassembly of Vipp1 oligomers
and thereby its membrane attachment. Binding to chloroplast
chaperones might also be an early step in Vipp1 import into
the organelle, to prevent unspecific aggregation of the protein
and to control oligomer formation (Fig. 4). Such an
assumption is supported by the recent observation that Vipp1
can be co-precipitated together with components of the TIC–
TOC protein import machinery of pea chloroplasts (Jouhet
and Gray, 2009). While a single Hsp70 and Hsp40 protein has
been identified in the stroma of Chlamydomonas chloroplasts
(Liu et al., 2005), multiple Hsp70/DnaK and Hsp40/DnaJ
proteins are encoded in cyanobacteria (Nimura et al., 2001;
Rupprecht et al., 2007, 2010; Sato et al., 2007; Duppre et al.,
2011). It is thus not clear yet which DnaK/DnaJ pair might
regulate assembly/disassembly of Vipp1 rings in cyanobacteria. However, involvement of a special cyanobacterial
Fig. 4. Import and assembly of Vipp1 in chloroplasts. Pre-Vipp1 is
imported from the cytoplasm via the translocons at the outer and
inner envelope of chloroplasts (TOC and TIC) into the chloroplast
stroma. Molecular chaperones prevent misfolding and unspecific
aggregation. Oligomerization of Vipp1 is guided by the Hsp70 and
Hsp40 chaperones in an ATP-dependent manner. Oligomeric
Vipp1 rings eventually interact with the inner envelope (IE) or
thylakoid membranes (TM).
DnaK and DnaJ protein in thylakoid membrane biogenesis
has been suggested (Nimura et al., 1996; Oguchi et al., 1997),
and thus it is intriguing to speculate that these proteins
interact with Vipp1. Such an assumption is supported by the
observation that the specified DnaK and DnaJ chaperones
are not encoded in the genome of the thylakoid-free
cyanobacterium Gloeobacter (Nakamura et al., 2003).
Hsp90C has also been reported to form a complex with
Hsp70, Hsp40, and Vipp1 in Clamydomonas chloroplasts
(Heide et al., 2009). It is noteworthy that an Arabidopsis
hsp90C mutant line has defects in chloroplast development
and thylakoid stacking (Cao et al., 2003). The point mutation
in Hsp90C in this mutant might affect interaction of
chloroplast Hsp90 with Vipp1 (Heide et al., 2009), which
subsequently perturbs thylakoid membrane biogenesis.
Involvement of Vipp1 in the integration of thylakoid
membrane proteins has been suggested based on the
observation of a direct interaction of Vipp1 with the
Albino3.2 protein in Chlamydomonas chloroplasts (Gohre
et al., 2006). The thylakoid-localized Albino3.2 belongs to
the conserved YidC/Oxa1p/Alb3 protein family, whose
members function as insertases, chaperones, and/or assembly
factors for transmembrane proteins (reviewed in Wang and
Dalbey, 2010). It has been suggested that Vipp1 is either
involved in membrane integration of Albino3.2 or that Vipp1
delivers substrates to Albino3.2 for membrane integration
(Gohre et al., 2006). Albino3.2 has an essential role during
assembly of photosystem II (PSII) and PSI reaction centres
in thylakoid membranes (Gohre et al., 2006). During
Albino3.2-mediated membrane integration and/or assembly
of protein complexes, the membrane structure might be
locally perturbed and Vipp1 could eventually be required to
stabilize the membrane. In Chlamydomonas, depletion of the
Albino3.2 protein results in overproduction of Vipp1 (Gohre
et al., 2006), while depletion of YidC in E. coli causes an
increased expression of PspA (van der Laan et al., 2003).
Thus, Vipp1 might be involved in stabilizing the membrane
structure during Albino3.2-mediated protein insertion but
also participates in maintaining the membrane integrity after
Albino3.2 depletion. Vipp1 (additionally?) could associate
with the TIC complex to stabilize the inner envelope
membrane in the vicinity of the TIC complex during TICmediated protein integration. Involvement of PspA-like
proteins in protein translocation/integration is further supported by the observation that increased PspA expression has
been observed in E. coli cells having a defect in the Sec
pathway (Kleerebezem and Tommassen, 1993). Furthermore,
overexpression of PspA in S. lividans improved the function
of both the Sec and Tat pathway (Vrancken et al., 2007).
PspA has been suggested to minimize proton leakage of the
bacterial plasma membrane during Tat-mediated protein
translocation/integration, and thus the occurrence of Vipp1
and/or PspA has been linked to the Tat pathway of bacteria,
archaea, and plant chloroplasts (DeLisa et al., 2004). Interestingly, this function can be taken over by the full-length
Vipp1 as well as by a C-terminally truncated Vipp1 of
Synechocystis PCC 6803 in an E. coli pspA deletion strain
(DeLisa et al., 2004).
Vipp1: a very import protein in plastids | 1707
Thylakoid membranes, the photosynthetic
electron transfer chain, and Vipp1
An increased amount of thylakoid membranes and chlorophylls correlates with increased light harvesting, and
chloroplasts adapt to high-light growth conditions by reduction of thylakoid membranes (Anderson et al., 1973;
Lichtenthaler et al., 1981; Melis and Harvey, 1981). Furthermore, in both chloroplasts and cyanobacteria, the ratio of
the two photosystems, PSI and PSII, alters in response to
changing light conditions (Chow et al., 1990; Kim et al.,
1993; Murakami et al., 1997). A comparative proteomic
analysis of bundle sheet and mesophyll chloroplasts in maize
has shown that mesophyll cell plastids, which contain a larger
thylakoid membrane system, have a 10-fold increased Vipp1
content (Majeran et al., 2005). Mesophyll cells also contain
high amounts of PSII, which is mostly lacking in bundle
sheet chloroplasts. Depletion of Vipp1 in both higher plants
and cyanobacteria resulted in a massive decrease of assembled thylakoid membrane layers and in defects in the
photosynthetic electron transfer chain (Kroll et al., 2001;
Westphal et al., 2001a). In Arabidopsis the defect could not
be attributed to a deficiency of a single photosynthetic
complex, but appeared to be caused by dysfunction of the
entire photosynthetic electron transfer chain (Kroll et al.,
2001). Vipp1-depleted plants displayed a dramatic reduction
of both photosystems as well as of the cytochrome b6f
complex. A disturbed energy transfer between antenna and
photosynthetic core complexes has also been noticed, and the
light-harvesting complexes are not properly attached to the
photosystems (Aseeva et al., 2007).
Electron microscopic analysis of Synechocystis Dvipp1
mutant cells showed a similar reduction in thylakoid
membranes. Furthermore, the overall thylakoid membrane
morphology was disturbed and the thylakoid membranes
were less well arranged than in the wild-type strain
(Fuhrmann et al., 2009b). This observation indicates that
the composition of the thylakoid membranes and/or thylakoid membrane modelling is affected by Vipp1 depletion. In
cyanobacteria, the two photosystems typically form dimers
and trimers, respectively, and trimerization of PSI was
affected by Vipp1 depletion (Fuhrmann et al., 2009b).
Furthermore, the relative PSI:PSII ratio was also significantly decreased in the depletion strain, which indicates an
impaired activity of the photosynthetic electron transfer
chain (Schneider et al., 2001, 2004; Volkmer et al., 2007).
After controlled depletion of Vipp1 in Synechocystis a drastic effect on photosynthetic complexes has been observed
(Gao and Xu, 2009), which further suggests a direct effect
of Vipp1 on photosystem biogenesis, assembly, or function.
Vipp1 and the need for vesicles in
chloroplasts and cyanobacteria
The debate on how thylakoid membranes are formed in
mature chloroplasts without a connection to the inner
envelope has been longstanding (reviewed in Vothknecht
and Westhoff, 2001). Several of the thylakoid constituents,
such as monogalactosidyl diacylglycerol and other lipids,
are synthesized within the inner envelope (Douce, 1974;
Joyard and Stumpf, 1981; Kobayashi et al., 2007). Consequently, there is a need for regulated transport between
internal membranes within chloroplasts. This might be
achieved by a vesicle transport system of eukaryotic origin
that was suggested to exist in chloroplasts of plants
(reviewed in von Wettstein, 2001). Vesicle formation can be
observed in chloroplasts of higher plants under certain
experimental conditions (Westphal et al., 2001b) but is no
longer detected in the Arabidopsis Vipp1 depletion strain
(Kroll et al., 2001). It is indeed conceivable that Vipp1 of
higher plants might support thylakoid biogenesis and
maintenance by sustaining vesicle traffic and thereby
ensuring the flow of lipids and other non-lipid components
from the inner envelope to the thylakoids.
It has long been debated whether thylakoid membranes
also represent a completely independent internal membrane
system in cyanobacteria or whether they are still linked to
the cytoplasmic membrane. However, recent structural
analyses strongly suggest that thylakoids indeed are not
directly connected to cyanobacterial cytoplasmic membranes, and thus represent a separate internal membrane
entity (Liberton et al., 2006; van de Meene et al., 2006;
Nevo et al., 2007; Schneider et al., 2007). Furthermore, it
has been suggested that precursors of the cyanobacterial
photosystems are pre-assembled within the plasma membrane (Zak et al., 2001; Keren et al., 2005), implying the
existence of a distinct lipid and protein transfer system from
the cytoplasmic to the thylakoid membrane. Such transfer
might involve vesicles or a temporary connection of the
membrane systems. Thylakoid centres in cyanobacteria may
be the origin of thylakoids, as these regions appear to
connect thylakoid membranes with the cytoplasmic membrane in some cyanobacteria (Nickelsen et al., 2010).
However, thylakoid centre structures are not typical and
highly conserved in these organisms, indicating that they are
not generally involved in thylakoid membrane biogenesis.
Vesicle accumulation was detected in chloroplasts of
A. thaliana and P. sativum by transmission electron
microscopy while providing conditions that are known to
prevent vesicle fusion in the cytosol. These include
a decrease in the temperature or treatment with toxins
such as ophiobolin A and microcystin, established inhibitors of cytosolic proteins involved in this process (Morre
et al., 1991; Kroll et al., 2001; Westphal et al., 2001b).
Until now, no vesicle transport system has been observed
under those conditions in cyanobacteria, green algae, or
even charophytes, implying that the specific vesicle transport system of chloroplasts might be restricted to
embryophytes (bryophytes, pteridophytes, and spermatophytes) and has evolved somewhere in an ancestor
between today’s charyophytes and bryophytes (Westphal
et al., 2003). Nevertheless, this has been a point of
ongoing debate, with different experimental evidence
supporting the occurrence of vesicle transport in photosynthetic organisms other than embryophytes (Hoober
1708 | Vothknecht et al.
et al., 1991). After staining the cyanobacterial cytoplasmic membrane with a fluorescent dye, intracellular
vesicular structures appeared within individual cells
(Schneider et al., 2007). Also, in the cyanobacterium
Microcoleus sp., large vesicular structures have been
identified by electron microscopy (Nevo et al., 2007).
Vipp1 might have a function similar to coat proteins in
the secretory pathway, which stabilize highly bent membranes during vesicle budding. For E. coli PspA it has been
observed that it predominantly binds to the cell poles of the
bacterium, where the membranes are highly bent and where
membrane stress and proton leakage might be higher than
in planar bilayer regions (Engl et al., 2009). Thus, Vipp1
might be involved in stabilizing highly bent membranes, as
is the case during vesicle formation. The diverse oligomeric
Vipp1 ring structures could also be involved in vesicle
constriction, similar to dynamins in eukaryotes (Fuhrmann
et al., 2009a). While assembly and disassembly of dynamin
is regulated by its GTPase domain, controlled disassembly
and formation of smaller Vipp1 rings eventually involves
Hsp70 and Hsp40, as discussed above. PspA and Vipp1
show homology to the eukaryotic Snf7 protein, which is
involved in protein sorting and transport from eukaryotic
endosomes to the vacuole/lysosome, and all three proteins
are clustered in a single protein family in the pfam database
(Finn et al., 2009). Furthermore, proteins with homologies to
components of the vesicular transfer system of the secretory
pathway have been predicted to localize to chloroplasts, and
some of the predicted proteins also have clear homologues in
cyanobacteria (Andersson and Sandelius, 2004). Thus, it
cannot be excluded per se that a vesicular transfer system
exists in chloroplasts and cyanobacteria.
Vipp1 function: does a common theme
emerge?
All studies carried out in chloroplasts and cyanobacteria
indicate that Vipp1 binds directly to membranes, locally
stabilizes bilayer structures, and thereby retains membrane
integrity. While initial observations indicated an involvement of Vipp1 in vesicle formation in chloroplasts, the
observed perturbations in an Arabidopsis vipp1 depletion
strain might be caused by an altered membrane structure
and/or composition. However, also during vesicle formation, membranes deform and Vipp1 might have a function
in stabilizing bent membranes, similar to coat proteins.
Involvement of Vipp1 in vesicle transfer between internal
membrane systems in chloroplasts and cyanobacteria is still
an appealing hypothesis as a general function of Vipp1,
since a membrane chaperone role could most probably also
be fulfilled by PspA, which is still present in cyanobacteria.
Nevertheless, Vipp1 has some distinct features different
from PspA. While both PspA and Vipp1 form ring
structures, the ring dimensions are dramatically different
and this leads to the necessity for variance in the PspA and
Vipp1 tertiary structure. PspA monomers most probably
form a kinked helix bundle, whereas Vipp1 may have a long
stretched a-helical shape (Bultema et al., 2010). These
different quaternary structures of PspA and Vipp1 eventually determine the exact physiological functions of the
individual proteins. A key to the specific function of Vipp1
could also be the C-terminal extension unique to Vipp1
proteins. This extension appears to correlate with the
appearance of thylakoid membranes, as Gloeobacter contains neither Vipp1 nor thylakoid membranes, whereas it
encodes a PspA protein, which could be crucially involved
in maintaining the integrity of the cytoplasmic membrane.
It is noteworthy that in the genome of Gloeobacter, genes
involved in sulphoquinovosyl diacylglycerol synthesis are
also missing (Nakamura et al., 2003). Sulphoquinovosyl
diacylglycerol is a constituent of typical thylakoid membranes, and, like phosphatidyl glycerol (PG), contains
a negatively charged head group. Depletion of Vipp1 in
Synechocystis results in a decreased PSI content and in
a reduced trimerization propensity of PSI (Fuhrmann et al.,
2009b), a phenotype also observed after PG depletion
(Hagio et al., 2000; Sato et al., 2000). Also in Arabidopsis
PG is required for photosynthesis, and a mutant having
inactivated PG synthesis was no longer able to grow
photoautotrophically and showed a severe reduction in
thylakoid membrane (Hagio et al., 2002). As PspA from
E. coli has been shown to interact preferentially with the
head groups of negatively charged lipids (Kobayashi et al.,
2007), all these observations link PspA and Vipp1 not only
generally to membranes, but more specifically to negatively
charged membrane lipids. Defined interaction of Vipp1 with
negatively charged lipids might thus enable spatial and
temporal control of Vipp1’s action on membrane systems in
cyanobacteria and chloroplasts.
However, all in all the exact function of both PspA and
Vipp1 remain elusive. Yet considering the importance of
Vipp1 for the viability of cyanobacteria and plants, it will
be essential in the future to obtain a more conclusive picture
of the role of this proteins in membrane biogenesis and
maintenance.
Acknowledgements
This work was supported by grants from the Deutsche
Forschungsgemeinschaft to UCV (SFB-TR1, project A6)
and DS (FOR 929, SCHN 690/3-1) and from the Center of
Complex Matter (COMATT) to DS. RH is supported by
a fellowship of the Max Planck Graduate Center Mainz.
References
Aldridge C, Cain P, Robinson C. 2009. Protein transport in
organelles: protein transport into and across the thylakoid membrane.
FEBS Journal 276, 1177–1186.
Anderson JM, Goodchild DJ, Boardman NK. 1973. Composition
of the photosystems and chloroplast structure in extreme shade
plants. Biochimica et Biophysica Acta 325, 573–585.
Vipp1: a very import protein in plastids | 1709
Andersson MX, Sandelius AS. 2004. A chloroplast-localized
vesicular transport system: a bio-informatics approach. BMC
Genomics 5, 40.
Duppre E, Rupprecht E, Schneider D. 2011. Specific and
promiscuous functions of multiple DnaJ proteins in Synechocystis sp.
PCC 6803. Microbiology 157, 1269–1278.
Aseeva E, Ossenbuhl F, Eichacker LA, Wanner G, Soll J,
Vothknecht UC. 2004. Complex formation of Vipp1 depends on its
alpha-helical PspA-like domain. Journal of Biological Chemistry 279,
35535–35541.
Elderkin S, Bordes P, Jones S, Rappas M, Buck M. 2005.
Molecular determinants for PspA-mediated repression of the AAA
transcriptional activator PspF. Journal of Bacteriology 187,
3238–3248.
Aseeva E, Ossenbuhl F, Sippel C, et al. 2007. Vipp1 is required for
basic thylakoid membrane formation but not for the assembly of
thylakoid protein complexes. Plant Physiology and Biochemistry 45,
119–128.
Elderkin S, Jones S, Schumacher J, Studholme D, Buck M.
2002. Mechanism of action of the Escherichia coli phage shock
protein PspA in repression of the AAA family transcription factor PspF.
Journal of Molecular Biology 320, 23–37.
Bergler H, Abraham D, Aschauer H, Turnowsky F. 1994. Inhibition
of lipid biosynthesis induces the expression of the pspA gene.
Microbiology 140, 1937–1944.
Engl C, Jovanovic G, Lloyd LJ, Murray H, Spitaler M, Ying L,
Errington J, Buck M. 2009. In vivo localizations of membrane stress
controllers PspA and PspG in Escherichia coli. Molecular Microbiology
73, 382–396.
Bidle KA, Kirkland PA, Nannen JL, Maupin-Furlow JA. 2008.
Proteomic analysis of Haloferax volcanii reveals salinity-mediated
regulation of the stress response protein PspA. Microbiology 154,
1436–1443.
Brissette JL, Russel M, Weiner L, Model P. 1990. Phage shock
protein, a stress protein of Escherichia coli. Proceedings of the
National Academy of Sciences, USA 87, 862–866.
Brissette JL, Weiner L, Ripmaster TL, Model P. 1991.
Characterization and sequence of the Escherichia coli stress-induced
psp operon. Journal of Molecular Biology 220, 35–48.
Bultema JB, Fuhrmann E, Boekema EJ, Schneider D. 2010.
Vipp1 and PspA: Related but not twins. Communicative and
Integrative Biology 3, 162–165.
Cao D, Froehlich JE, Zhang H, Cheng CL. 2003. The chlorateresistant and photomorphogenesis-defective mutant cr88 encodes
a chloroplast-targeted HSP90. The Plant Journal 33, 107–118.
Chow WS, Melis A, Anderson JM. 1990. Adjustments of
photosystem stoichiometry in chloroplasts improve the quantum
efficiency of photosynthesis. Proceedings of the National Academy of
Sciences, USA 87, 7502–7506.
Crooks GE, Hon G, Chandonia JM, Brenner SE. 2004. WebLogo:
a sequence logo generator. Genome Research 14, 1188–1190.
Cserzo M, Wallin E, Simon I, von Heijne G, Elofsson A. 1997.
Prediction of transmembrane alpha-helices in prokaryotic membrane
proteins: the dense alignment surface method. Protein Engineering 10,
673–676.
Darwin AJ. 2005. The phage-shock-protein response. Molecular
Microbiology 57, 621–628.
Darwin AJ, Miller VL. 2001. The psp locus of Yersinia enterocolitica
is required for virulence and for growth in vitro when the Ysc type III
secretion system is produced. Molecular Microbiology 39, 429–444.
DeLisa MP, Lee P, Palmer T, Georgiou G. 2004. Phage shock
protein PspA of Escherichia coli relieves saturation of protein export via
the Tat pathway. Journal of Bacteriology 186, 366–373.
Douce R. 1974. Site of biosynthesis of galactolipids in spinach
chloroplasts. Science 183, 852–853.
Dufresne A, Salanoubat M, Partensky F, et al. 2003. Genome
sequence of the cyanobacterium Prochlorococcus marinus SS120,
a nearly minimal oxyphototrophic genome. Proceedings of the
National Academy of Sciences, USA 100, 10020–10025.
Eriksson S, Lucchini S, Thompson A, Rhen M, Hinton JC. 2003.
Unravelling the biology of macrophage infection by gene expression
profiling of intracellular Salmonella enterica. Molecular Microbiology 47,
103–118.
Finn RD, Mistry J, Tate J, et al. 2009. The Pfam protein families
database. Nucleic Acids Research 38, D211–D222.
Fuhrmann E, Bultema JB, Kahmann U, Rupprecht E,
Boekema EJ, Schneider D. 2009a. The vesicle-inducing protein 1
from Synechocystis sp. PCC 6803 organizes into diverse higherordered ring structures. Molecular Biology of the Cell 20,
4620–4628.
Fuhrmann E, Gathmann S, Rupprecht E, Golecki J, Schneider D.
2009b. Thylakoid membrane reduction affects the photosystem
stoichiometry in the cyanobacterium Synechocystis sp. PCC 6803.
Plant Physiology 149, 735–744.
Gao H, Xu X. 2009. Depletion of Vipp1 in Synechocystis sp. PCC
6803 affects photosynthetic activity before the loss of thylakoid
membranes. FEMS Microbiol Letters 292, 63–70.
Gohre V, Ossenbuhl F, Crevecoeur M, Eichacker LA,
Rochaix JD. 2006. One of two alb3 proteins is essential for the
assembly of the photosystems and for cell survival in Chlamydomonas.
The Plant Cell 18, 1454–1466.
Guindon S, Gascuel O. 2003. A simple, fast, and accurate algorithm
to estimate large phylogenies by maximum likelihood. Systematic
Biology 52, 696–704.
Hagio M, Gombos Z, Varkonyi Z, Masamoto K, Sato N,
Tsuzuki M, Wada H. 2000. Direct evidence for requirement of
phosphatidylglycerol in photosystem II of photosynthesis. Plant
Physiology 124, 795–804.
Hagio M, Sakurai I, Sato S, Kato T, Tabata S, Wada H. 2002.
Phosphatidylglycerol is essential for the development of thylakoid
membranes in Arabidopsis thaliana. Plant and Cell Physiology 43,
1456–1464.
Hankamer BD, Elderkin SL, Buck M, Nield J. 2004. Organization
of the AAA(+) adaptor protein PspA is an oligomeric ring. Journal of
Biological Chemistry 279, 8862–8866.
Heide H, Nordhues A, Drepper F, Nick S, Schulz-Raffelt M,
Haehnel W, Schroda M. 2009. Application of quantitative
immunoprecipitation combined with knockdown and cross-linking to
1710 | Vothknecht et al.
Chlamydomonas reveals the presence of vesicle-inducing protein in
plastids 1 in a common complex with chloroplast HSP90C.
Proteomics 9, 3079–3089.
Kleerebezem M, Tommassen J. 1993. Expression of the pspA gene
stimulates efficient protein export in Escherichia coli. Molecular
Microbiology 7, 947–956.
Honda D, Yokota A, Sugiyama J. 1999. Detection of seven major
evolutionary lineages in cyanobacteria based on the 16S rRNA gene
sequence analysis with new sequences of five marine Synechococcus
strains. Journal of Molecular Evolution 48, 723–739.
Kobayashi H, Yamamoto M, Aono R. 1998. Appearance of
a stress-response protein, phage-shock protein A, in Escherichia coli
exposed to hydrophobic organic solvents. Microbiology 144, 353–359.
Hoober JK, Boyd CO, Paavola LG. 1991. Origin of thylakoid
membranes in Chlamydomonas reinhardtii Y-1 at 38C. Plant
Physiology 96, 1321–1328.
Huang F, Fulda S, Hagemann M, Norling B. 2006. Proteomic
screening of salt-stress-induced changes in plasma membranes of
Synechocystis sp. strain PCC 6803. Proteomics 6, 910–920.
Hyyrylainen HL, Sarvas M, Kontinen VP. 2005. Transcriptome
analysis of the secretion stress response of Bacillus subtilis. Applied
Microbiology and Biotechnology 67, 389–396.
Im CS, Grossman AR. 2002. Identification and regulation of high
light-induced genes in Chlamydomonas reinhardtii. The Plant Journal
30, 301–313.
Joly N, Burrows PC, Engl C, Jovanovic G, Buck M. 2009. A lowerorder oligomer form of phage shock protein A (PspA) stably associates
with the hexameric AAA(+) transcription activator protein PspF for
negative regulation. Journal of Molecular Biology 394, 764–775.
Joly N, Engl C, Jovanovic G, Huvet M, Toni T, Sheng X,
Stumpf MP, Buck M. 2010. Managing membrane stress: the phage
shock protein (Psp) response, from molecular mechanisms to
physiology. FEMS Microbiology Reviews 34, 797–827.
Jouhet J, Gray JC. 2009. Interaction of actin and the chloroplast
protein import apparatus. Journal of Biological Chemistry 284,
19132–19141.
Jovanovic G, Engl C, Buck M. 2009. Physical, functional and
conditional interactions between ArcAB and phage shock proteins
upon secretin-induced stress in Escherichia coli. Molecular
Microbiology 74, 16–28.
Jovanovic G, Lloyd LJ, Stumpf MP, Mayhew AJ, Buck M. 2006.
Induction and function of the phage shock protein extracytoplasmic
stress response in Escherichia coli. Journal of Biological Chemistry
281, 21147–21161.
Joyard J, Stumpf PK. 1981. Synthesis of long-chain Acyl-CoA in
chloroplast envelope membranes. Plant Physiology 67, 250–256.
Kaneko T, Nakamura Y, Wolk CP, et al. 2001. Complete genomic
sequence of the filamentous nitrogen-fixing cyanobacterium Anabaena
sp. strain PCC 7120. DNA Research 8, 205–213; 227–253.
Keren N, Liberton M, Pakrasi HB. 2005. Photochemical
competence of assembled photosystem II core complex in
cyanobacterial plasma membrane. Journal of Biological Chemistry
280, 6548–6553.
Kobayashi K, Kondo M, Fukuda H, Nishimura M, Ohta H. 2007.
Galactolipid synthesis in chloroplast inner envelope is essential for proper
thylakoid biogenesis, photosynthesis, and embryogenesis. Proceedings
of the National Academy of Sciences, USA 104, 17216–17221.
Kobayashi R, Suzuki T, Yoshida M. 2007. Escherichia coli phageshock protein A (PspA) binds to membrane phospholipids and repairs
proton leakage of the damaged membranes. Molecular Microbiology
66, 100–109.
Kroll D, Meierhoff K, Bechtold N, Kinoshita M, Westphal S,
Vothknecht UC, Soll J, Westhoff P. 2001. VIPP1, a nuclear gene of
Arabidopsis thaliana essential for thylakoid membrane formation.
Proceedings of the National Academy of Sciences, USA 98,
4238–4242.
Lawrence ME, Possingham JV. 1984. Observations of microtubulelike structures within spinach plastids. Colchester, UK: Portland.
Li HM, Kaneko Y, Keegstra K. 1994. Molecular cloning of
a chloroplastic protein associated with both the envelope and
thylakoid membranes. Plant Molecular Biology 25, 619–632.
Liberton M, Howard Berg R, Heuser J, Roth R, Pakrasi HB.
2006. Ultrastructure of the membrane systems in the unicellular
cyanobacterium Synechocystis sp. strain PCC 6803. Protoplasma
227, 129–138.
Lichtenthaler H, Buschmann C, Döll M, Fietz HJ, Bach T,
Kozel U, Meier D, Rahmsdorf U. 1981. Photosynthetic activity,
chloroplast ultrastructure, and leaf characteristics of high-light and
low-light plants and of sun and shade leaves. Photosynthesis
Research 2, 115–141.
Liu C, Willmund F, Golecki JR, Cacace S, Hess B, Markert C,
Schroda M. 2007. The chloroplast HSP70B-CDJ2-CGE1 chaperones
catalyse assembly and disassembly of VIPP1 oligomers in
Chlamydomonas. The Plant Journal 50, 265–277.
Liu C, Willmund F, Whitelegge JP, Hawat S, Knapp B, Lodha M,
Schroda M. 2005. J-domain protein CDJ2 and HSP70B are
a plastidic chaperone pair that interacts with vesicle-inducing protein in
plastids 1. Molecular Biology of the Cell 16, 1165–1177.
Lloyd LJ, Jones SE, Jovanovic G, Gyaneshwar P, Rolfe MD,
Thompson A, Hinton JC, Buck M. 2004. Identification of a new
member of the phage shock protein response in Escherichia coli, the
phage shock protein G (PspG). Journal of Biological Chemistry 279,
55707–55714.
Kim JH, Glick RE, Melis A. 1993. Dynamics of photosystem
stoichiometry adjustment by light quality in chloroplasts. Plant
Physiology 102, 181–190.
Lucchini S, Liu H, Jin Q, Hinton JC, Yu J. 2005. Transcriptional
adaptation of Shigella flexneri during infection of macrophages and
epithelial cells: insights into the strategies of a cytosolic bacterial
pathogen. Infection and Immunity 73, 88–102.
Kleerebezem M, Crielaard W, Tommassen J. 1996. Involvement of
stress protein PspA (phage shock protein A) of Escherichia coli in
maintenance of the protonmotive force under stress conditions. EMBO
Journal 15, 162–171.
Majeran W, Cai Y, Sun Q, van Wijk KJ. 2005. Functional
differentiation of bundle sheath and mesophyll maize chloroplasts
determined by comparative proteomics. The Plant Cell 17,
3111–3140.
Vipp1: a very import protein in plastids | 1711
Mascher T, Zimmer SL, Smith TA, Helmann JD. 2004. Antibioticinducible promoter regulated by the cell envelope stress-sensing twocomponent system LiaRS of Bacillus subtilis. Antimicrobial Agents and
Chemotherapy 48, 2888–2896.
Rupprecht E, Duppre E, Schneider D. 2010. Similarities and
singularities of three DnaK proteins from the cyanobacterium
Synechocystis sp. PCC 6803. Plant and Cell Physiology 51,
1210–1218.
Melis A, Harvey GW. 1981. Regulation of photosystem
stoichiometry, chlorophyll a and chlorophyll b content and relation to
chloroplast ultrastructure. Biochimica et Biophysica Acta 637,
138–145.
Rupprecht E, Gathmann S, Fuhrmann E, Schneider D. 2007.
Three different DnaK proteins are functionally expressed in the
cyanobacterium Synechocystis sp. PCC 6803. Microbiology 153,
1828–1841.
Mimuro M, Tomo T, Tsuchiya T. 2008. Two unique cyanobacteria
lead to a traceable approach of the first appearance of oxygenic
photosynthesis. Photosynthesis Research 97, 167–176.
Sato M, Nimura-Matsune K, Watanabe S, Chibazakura T,
Yoshikawa H. 2007. Expression analysis of multiple dnaK genes in
the cyanobacterium Synechococcus elongatus PCC 7942. Journal of
Bacteriology 189, 3751–3758.
Morre DJ, Sellden G, Sundqvist C, Sandelius AS. 1991. Stromal
low temperature compartment derived from the inner membrane of the
chloroplast envelope. Plant Physiology 97, 1558–1564.
Murakami A, Kim SJ, Fujita Y. 1997. Changes in photosystem
stoichiometry in response to environmental conditions for cell growth
observed with the cyanophyte Synechocystis PCC 6714. Plant and
Cell Physiology 38, 392–397.
Nakamura Y, Kaneko T, Sato S, et al. 2003. Complete genome
structure of Gloeobacter violaceus PCC 7421, a cyanobacterium that
lacks thylakoids. DNA Research 10, 137–145.
Nelissen B, Van de Peer Y, Wilmotte A, De Wachter R. 1995. An
early origin of plastids within the cyanobacterial divergence is
suggested by evolutionary trees based on complete 16S rRNA
sequences. Molecular Biology and Evolution 12, 1166–1173.
Nevo R, Charuvi D, Shimoni E, Schwarz R, Kaplan A, Ohad I,
Reich Z. 2007. Thylakoid membrane perforations and connectivity
enable intracellular traffic in cyanobacteria. EMBO Journal 26,
1467–1473.
Nickelsen J, Rengstl B, Stengel A, Schottkowski M, Soll J,
Ankele E. 2010. Biogenesis of the cyanobacterial thylakoid
membrane system—an update. FEMS Microbiology Letters 315, 1–5.
Nimura K, Takahashi H, Yoshikawa H. 2001. Characterization of
the dnaK multigene family in the Cyanobacterium Synechococcus sp.
strain PCC7942. Journal of Bacteriology 183, 1320–1328.
Nimura K, Yoshikawa H, Takahashi H. 1996. DnaK3, one of the
three DnaK proteins of cyanobacterium Synechococcus sp.
PCC7942, is quantitatively detected in the thylakoid membrane.
Biochemical and Biophysical Research Communications 229,
334–340.
Oguchi K, Nimura K, Yoshikawa H, Takahashi H. 1997. Sequence
and analysis of a dnaJ homologue gene in cyanobacterium
Synechococcus sp. PCC7942. Biochemical and Biophysical Research
Communications 236, 461–466.
Petersohn A, Brigulla M, Haas S, Hoheisel JD, Volker U,
Hecker M. 2001. Global analysis of the general stress response of
Bacillus subtilis. Journal of Bacteriology 183, 5617–5631.
Pietiainen M, Gardemeister M, Mecklin M, Leskela S, Sarvas M,
Kontinen VP. 2005. Cationic antimicrobial peptides elicit a complex
stress response in Bacillus subtilis that involves ECF-type sigma
factors and two-component signal transduction systems. Microbiology
151, 1577–1592.
Rippka R, Waterbury J, Cohen-Bazire G. 1974. A cyanobacterium
which lacks thylakoids. Archives of Microbiology 100, 419–436.
Sato N, Hagio M, Wada H, Tsuzuki M. 2000. Requirement of
phosphatidylglycerol for photosynthetic function in thylakoid
membranes. Proceedings of the National Academy of Sciences, USA
97, 10655–10660.
Savva CG, Dewey JS, Deaton J, White RL, Struck DK,
Holzenburg A, Young R. 2008. The holin of bacteriophage lambda
forms rings with large diameter. Molecular Microbiology 69, 784–793.
Schneider D, Berry S, Rich P, Seidler A, Rogner M. 2001. A
regulatory role of the PetM subunit in a cyanobacterial cytochrome b6f
complex. Journal of Biological Chemistry 276, 16780–16785.
Schneider D, Berry S, Volkmer T, Seidler A, Rogner M. 2004.
PetC1 is the major Rieske iron–sulfur protein in the cytochrome b6f
complex of Synechocystis sp. PCC 6803. Journal of Biological
Chemistry 279, 39383–39388.
Schneider D, Fuhrmann E, Scholz I, Hess WR, Graumann PL.
2007. Fluorescence staining of live cyanobacterial cells suggest nonstringent chromosome segregation and absence of a connection
between cytoplasmic and thylakoid membranes. BMC Cell Biology 8,
39.
Schnell DJ, Blobel G, Keegstra K, Kessler F, Ko K, Soll J. 1997.
A consensus nomenclature for the protein-import components of the
chloroplast envelope. Trends in Cell Biology 7, 303–304.
Srivastava R, Pisareva T, Norling B. 2005. Proteomic studies of the
thylakoid membrane of Synechocystis sp. PCC 6803. Proteomics 5,
4905–4916.
Standar K, Mehner D, Osadnik H, Berthelmann F, Hause G,
Lunsdorf H, Bruser T. 2008. PspA can form large scaffolds in
Escherichia coli. FEBS Letters 582, 3585–3589.
Suzuki I, Kanesaki Y, Mikami K, Kanehisa M, Murata N. 2001.
Cold-regulated genes under control of the cold sensor Hik33 in
Synechocystis. Molecular Microbiology 40, 235–244.
Thompson JD, Gibson TJ, Plewniak F, Jeanmougin F,
Higgins DG. 1997. The CLUSTAL_X windows interface: flexible
strategies for multiple sequence alignment aided by quality analysis
tools. Nucleic Acids Research 25, 4876–4882.
van de Meene AM, Hohmann-Marriott MF, Vermaas WF,
Roberson RW. 2006. The three-dimensional structure of the
cyanobacterium Synechocystis sp. PCC 6803. Archives of
Microbiology 184, 259–270.
van der Laan M, Urbanus ML, Ten Hagen-Jongman CM,
Nouwen N, Oudega B, Harms N, Driessen AJ, Luirink J. 2003. A
conserved function of YidC in the biogenesis of respiratory chain
1712 | Vothknecht et al.
complexes. Proceedings of the National Academy of Sciences, USA
Wang P, Dalbey RE. 2010. Inserting membrane proteins: the YidC/
100, 5801–5806.
Oxa1/Alb3 machinery in bacteria, mitochondria, and chloroplasts.
Volkmer T, Schneider D, Bernat G, Kirchhoff H, Wenk SO,
Biochimica et Biophysica Acta 1808, 866–875.
Rogner M. 2007. Ssr2998 of Synechocystis sp. PCC 6803 is involved
Weiner L, Model P. 1994. Role of an Escherichia coli stress-
in regulation of cyanobacterial electron transport and associated with
response operon in stationary-phase survival. Proceedings of the
the cytochrome b6f complex. Journal of Biological Chemistry 282,
National Academy of Sciences, USA 91, 2191–2195.
3730–3737.
Westphal S, Heins L, Soll J, Vothknecht UC. 2001a. Vipp1
von Wettstein D. 2001. Discovery of a protein required for
deletion mutant of Synechocystis: a connection between bacterial
photosynthetic membrane assembly. Proceedings of the National
phage shock and thylakoid biogenesis? Proceedings of the National
Academy of Sciences, USA 98, 3633–3635.
Academy of Sciences, USA 98, 4243–4248.
Vothknecht UC, Westhoff P. 2001. Biogenesis and origin of
Westphal S, Soll J, Vothknecht UC. 2001b. A vesicle transport
thylakoid membranes. Biochimica et Biophysica Acta 1541,
system inside chloroplasts. FEBS Letters 506, 257–261.
91–101.
Westphal S, Soll J, Vothknecht UC. 2003. Evolution of chloroplast
Vrancken K, De Keersmaeker S, Geukens N, Lammertyn E,
vesicle transport. Plant and Cell Physiology 44, 217–222.
Anne J, Van Mellaert L. 2007. pspA overexpression in
Wolf D, Kalamorz F, Wecke T, et al. 2010. In-depth profiling of the
Streptomyces lividans improves both Sec- and Tat-dependent protein
LiaR response of Bacillus subtilis. Journal of Bacteriology 192,
secretion. Applied Microbiology and Biotechnology 73,
4680–4693.
1150–1157.
Zak E, Norling B, Maitra R, Huang F, Andersson B, Pakrasi HB.
Vrancken K, Van Mellaert L, Anne J. 2008. Characterization of the
2001. The initial steps of biogenesis of cyanobacterial photosystems
Streptomyces lividans PspA response. Journal of Bacteriology 190,
occur in plasma membranes. Proceedings of the National Academy of
3475–3481.
Sciences, USA 98, 13443–13448.