Bioscience Reports, Vol. 16, No. 2, 1996
REVIEW
A Current Assessment of Photosystem II
Structure
William V. Nichoison, la Robert C. Ford, TM and Andreas
Holzenburg 3
Received July 14, 1995; accepted November 24, 1995
This review covers the recent progress in the elucidation of the structure of photosystem lI (PSII).
Because much of the structural information for this membrane protein complex has been revealed by
electron microscopy (EM), the review will also consider the specific technical and interpretation
problems that arise with EM where they are of particular relevance to the structural data. Most recent
reviews of photosystem II structure have concentrated on molecular studies of the PSII genes and on
the likely roles of the subunits that they encode or they were mainly concerned with the biophysical
data and fast absorption spectroscopy largely relating to electron transfer in various purified PSII
preparations. In this review, we will focus on the approaches to the three-dimensional architecture of
the complex and the lipid bilayer in which it is located (the thylakoid membrane) with special
emphasis placed upon electron microscopical studies of PSII-containing thylakoid membranes. There
are a few reports of 3D crystals of PSII and of associated X-ray diffraction measurements and
although little structural information has so far been obtained from such studies (because of the lack
of 3D crystals of sufficient quality), the prospects for such studies are also assessed.
KEY WORDS: Electron microscopy; photosystem II; thylakoid membrane.
A B B R E V I A T I O N S : ATP, adenosine triphosphate; Chl, chlorophyll; CP, chlorophyll-binding protein;
EM, electron microscopy; LHC, light harvesting complex; NADP, nicotinamide adenine dinucleotide
phosphate; OEC, oxygen evolution enhancing complex; PS, photosystem; Tris, tris-hydroxymethyl
aminomethane.
THE BIOLOGICAL ROLE A N D LOCATION OF PHOTOSYSTEM H
Photosystem II is the oxygen-evolving enzyme involved in photosynthesis. It
catalyses the oxidation of water using light energy. The electrons from the
oxidation are used to reduce plastoquinone initially, and are then passed along
1Department of Biochemistry and Applied Molecular Biology, UMIST, PO Box 88, Manchester
M60 1QD, UK.
2 Present address: Department of Chemistry, University of Glasgow, Glasgow G12 8QQ, UK.
3 Department of Biochemistry and Molecular Biology and Department of Genetics, University of
Leeds, Leeds LS2 9JT, UK.
4 To whom correspondence should be addressed.
159
0144-8463/96/0400-0159509.50/09 1996PlenumPublishingCorporation
i60
Nicholson, Ford, and Holzenburg
stroma
.,;-//
'~::.
~
0
~ " - ~ "~.
~
,~"
i
-- ~.
~s ~
f,,/-)
..,-
.....
lumen
stroma (partition region)
r.-t
LHCII ~
"
Fig. la. Simplified representation of a higher plant chloroplast and the
location of PSII complexes in the grana thylakoid membranes (zoomed
area). The internal membranes of the chloroplast (thylakoids) are arranged
as inter-connected grana stacks and non-appressed lamellae (NAL) which
separate the chloroplast into two aqueous compartments, the stroma and the
lumen. PSII is thought to be located primarily in the grana whilst PSI and
the H+-ATPase are thought to be located in the non-appressed lamellae,
with an intermediate distribution predicted for the cytochrome bJf complex.
PSII complexes in separate stacked membranes approach closely on the
stromal side of the membrane. This so-called partition region is only a few
nm across. To give an idea of scale, the PSII complex is about 9 nm across
with a 4.5 nm thick lipid bilayer. The chloroplast is usually 2-10 micometres
across, with grana stacks about 500 nm in diameter.
an electron transfer chain (via the c y t o c h r o m e b6f c o m p l e x and p h o t o s y s t e m I) to
eventually be used to r e d u c e N A D P . T h e p r o t o n s transported across the
thylakoid m e m b r a n e , establish an electrochemical gradient which is used to
g e n e r a t e A T P (the c y t o c h r o m e b6f complex and p h o t o s y s t e m I are also involved
in generating the electrochemical gradient across the thylakoid m e m b r a n e ) .
P h o t o s y s t e m II is a m e m b r a n e protein c o m p l e x and is f o u n d in the thylakoid
m e m b r a n e s of chloroplasts (in plants) or c y a n o b a c t e r i a (prokaryotes). A schematic view of higher plant P S I I and its location in the chloroplast is shown in Fig. la.
PSII PURIFICATION A N D POLYPEPTIDE COMPOSITION
Isolation of P S I I - e n r i c h e d grana m e m b r a n e s is possible using higher plant
thylakoid m e m b r a n e s [Berthold et al. (1981); F o r d & E v a n s (1983)]. Solubilisation of P S I I with very mild detergents such as d o d e c y l maltoside or octyl
PhotosystemII
161
gtucoside at low detergent: chlorophyll ratios (<10:lwt/wt) gives rise to a
soluble PSII complex that can be purified by conventional protein purification
methods [Ghanotakis et al. (1989); Ghanotakis & Yocum (1990); Vermaas,
(1993)]. Purification involving more extensive detergent exposure usually causes
the dissociation of part of the complex, especially the group of proteins known as
the peripheral light-harvesting antennae proteins. A residual core complex of
PSII remains intact in most of the currently employed purification procedures.
The core can be prepared with water-splitting activity preserved and it also
retains some of the extrinsic polypeptides associated with stimulating oxygen
evolution (Haag et al. (1990); Bricker et al. (1985); Tang & Satoh (1985); Y u a s a et
al. (1984)]. A harsher treatment of the purified core complex by extensive
washing with detergent can be used to separate PSII core components from each
other. Two further light-harvesting proteins termed CP47 and CP43 can be
isolated as well as a reaction centre of PSII [Nanba & Satoh (1987); Chapman et
al. (1991); Fotinou & Ghanotakis (1990)]. It has not yet been possible to prepare
the reaction centre with any preservation of the water-splitting activity, but it
retains light-induced electron transfer activity. The reaction centre can be
dissociated under harsh conditions and is composed of at least four polypeptides.
Higher plant PSII core complex is known to be composed of several integral
membrane proteins [Rutherford, (1989); Ghanotakis & Yocum (1990)]. These are
termed D1 (a 32 kDa polypeptide); D2 (34 kDa); two cytochromes b559 (each
composed of a 6 kDa and a 9 kDa polypeptide); and the two chlorophyll binding
polypeptides CP43 and CP47 (with apparent molecular masses of 43 kDa and
47 kDa, respectively). Also associated with the core are three extrinsic proteins of
17 kDa, 23 kDa and 33 kDa. Other polypeptides have been observed in some (but
not all) PSII core preparations by using gel electrophoresis techniques which
allow good resolution of polypeptides with low molecular mass [(Hansson &
Wydrzynski (1990); Ikeuchi (1992); Jansson et al. (1992)]. These include PSII-S,
PSII-R, PSII-H, PSII-T, PSII-L and PSII-K of apparent molecular masses in the
range 22 to 4 kDa [Ljungberg et al. (1986); Lautner et al. (1988); Ikeuchi & Inoue
(1988)]. Functions have not yet been identified for these polypeptides although it
is now generally accepted that they are all subunits of PSII.
D1 and D2 are homologous polypeptides and form a heterodimer. The
D1/D2 complex is the location of the reaction centre electron transfer components which include chlorophylls, pheophytins and quinones [Tang et al.,
(1990)]. The function of cytochrome b559 in the complex is less clear. It has been
suggested that it may have a protective role [Thompson & Brudvig (1988)]. The
two chlorophyll binding polypeptides, CP47 and CP43, are integral lightharvesting polypeptides in the complex and bind chlorophyll a and carotenoids
[Hansson & Wydrzynski (1990)]. They also have somewhat similar amino acid
sequences and both are characterised by large extramembraneous loops that are
exposed on the lumenal side of the complex. The three extrinsic polypeptides
were initially thought to be the site of the Mn cluster as experiments showed that
they were required for oxygen-evolving activity [Murata & Miyao (1985)].
However, each of them is functionally replacable by Ca 2+ and/or C1- ions.
Together, the 3 polypeptides form the oxygen-evolving complex (OEC) or
162
Nicholson, Ford, and Holzenburg
oxygen evolution enhancing complex. They may have a role in stabilising the Mn
cluster, but it is unlikely that they directly contribute to Mn co-ordination.
Cyanobacterial PSII lacks the 23 and 17 kDa OEC polypeptides and shows little
sensitivity to Ca 2+ or C1- ion concentration provided that the 33 kDa protein is
present [Ikeuchi (1992); Satoh & Katoh (1985); Ohno et al. (1986)].
The higher plant light-harvesting antennae contains a number of related
polypeptides, some of which have only been identified recently [Ikeuchi (1992)].
T h e y are polypeptides that bind both chlorophyll a and b as well as carotenoids,
with apparent molecular masses for the apoproteins of about 25-30 kDa. The
most abundant of the light-harvesting polypeptides is light-harvesting complex II
or LHCII (also sometimes called LHCP). The 3D structure of isolated LHCII was
obtained to high resolution by electron crystallography [Kuhlbrandt & Wang
(1991), Kuhlbrandt et al., (1994)]. LHCII is trimeric in the crystallised form and
current thinking has it to be present in vivo also as a trimer. However, other
related light-harvesting polypeptides appear to be monomeric [Jannson (1994)].
Cyanobacterial PSII lacks the peripheral Chl a / b binding polypeptides; (the
phycobiliproteins, organised into large macromolecular pigment-protein complexes are thought to perform this light-harvesting function in cyanobacteria)
[Hansson & Wydrzynski (1990)].
The picture that emerges from these various biochemical extraction procedures and polypeptide analyses is a PSII complex that consists of a series of 3
shells: (i) An easily removed and variable peripheral core of light-harvesting
chlorophyll a / b proteins. (ii) A more stable and constant core of Chl a binding
proteins (CP47 and CP43). (iii) An inner core containing the final trap of
excitation energy (the P680 Chl a dimer) and associated electron transfer
components bound by a heterodimer of two integral membrane proteins. In
cyanobacteria, the PSII outer shell (i) is not present (this group of photosynthetic
bacteria do not produce chlorophyll b), but is replaced by an extrinsic (water
soluble) light-harvesting complex (the phycobilisome) that can be readily washed
off the membrane surface.
A comparison of the various plant and cyanobacterial core complex
preparations that have been reported suggests that they contain 40 to 80 Chl a
molecules for every PSII electron transfer chain [Ghanotakis & Yocum (1986),
Enami et al., (1989); Haag et al., (1990); van Leeuwen et al., (1991); de Vitry et
al., (1991); Roegner et al., (1987); Brettel et al., (1985)]. Since 200 to 300 Chl a
and b molecules are associated with every PSII electron transfer chain in plant
thylakoids and PSII-enriched grana membranes [Ford & Evans (1983), Brettel et
al., (1985)], this means that between 120 and 260 Chl a and b molecules are
non-core in PSII. This estimate has two areas of relevance for this review. Firstly
it shows the potential for dynamic modification of light harvesting in plant PSII,
since a fair proportion of these peripheral chlorophylls can reversibly associate
with PSII in response to varying light conditions in vivo (in cyanobacteria, a
similar dynamic adaptation mechanism exists for the phycobilisome). Secondly,
using the figures we can estimate the number of light-harvesting proteins (such as
LHCII and CP29) which will be associated with each plant PSII, since it is known
that 11 to 15 Chl a and b molecules are associated with each polypeptide. Thus,
PhotosystemII
163
at least 8 and at most 20LHC-type polypeptides are associated with each PSII
unit. This number has great significance for analysis of the size of the complex
(see later).
X - R A Y C R Y S T A L L O G R A P H I C STUDIES
As yet, no X-ray structure of PSII has been produced, althought structures
for light-harvesting subunits of higher plant and cyanobacterial PSII have been
obtained by electron and X-ray crystallography respectively [Kuhlbrandt et al.,
(1994); Schirmer et aL, (1986)]. Small 3D crystals of the spinach PSII core
complex have been obtained [Fotinou et al. (1993); Adir et al. (1992)]. The latter
group obtained crystals which diffracted to about 15 A resolution with their
hexagonal crystals; the former group obtained diffraction orders to about 10]k
resolution, the crystals consist of CP47, CP43, cytochrome b559, D1, D2 and 3
small polypeptides (of less than 10 kDa). The pruified PSII core complex, from
which the crystals were obtained, retained reaction centre activity but not oxygen
evolution activity. Unit cell dimensions were not calculated for either case,
although Fotinou et al. (1993) estimate unit cell axes to be about 200 A, which
seems reasonable given the large size of the core complex. Crystals of PSII have
been obtained from the thermophilic cyanobacterium Mastigocladus laminosus
[Adir et al. (1994)]. These crystals diffracted X-rays to 2.9 A resolution, however,
the unusual unit cell dimensions of 160 • 42 • 160 A, (i.e. too small along one
axis of the unit ceil to be compatible with PSII), suggest that some other
cyanobacterial protein was crystallised. In their report, the authors stated that
they had still to investigate the polypeptide composition of their crystals by gel
electrophoresis.
H O M O L O G Y STUDIES
The
structure
of
the
reaction
centres
of
the
purple
bacteria
R h o d o p s e u d o m o n a s viridis [Deisenhofer et al. (1984); Deisenhofer et al. (1985);
Michel et al. (1986) and Rhodobacter sphaeroides have been obtained [Allen et al.
(1987); Yeates et aL (1987)], and from this data it was inferred that the D1 and
D2 subunits of PSII probably have structural similarities with the L and M
subunits of the purple bacteria reaction centre. There are specific sequence
homologies between the L subunit and the D1 subunit and both can be labelled
by azidoatrazine which binds at the herbicide or Qb binding site of D1 [Pfister
(1981)]. Hence D1 was proposed to correspond to the L subunit and D2 to the M
subunit [Michel & Deisenhofer (1988); Hannson & Wydrzynski (1990)]. The
co-factors of the electron acceptor side and the primary electron donor (the P680
chlorophyll dimer) of the PSII reaction centre are similar to those in the purple
bacterial reaction centre. In most models of the structure they are located on the
D1 and D2 polypeptides. Of these co-factors, P680, pheophytin, Qa and Qb have
been observed spectroscopically [Rutherford (1989)]. 2 extra pheophytin and 2
164
Nicholson,Ford, and Holzenburg
extra chlorophylls are placed in models of PSII structure;" as they appear in the
crystallographic structures of the purple bacterial reaction centre [Ruffle et al.,
(1992)]. The electron donor side of PSII differs from the purple bacterial reaction
centre. A tyrosine radical, Tyr Z is the electron donor to P680 and is observed in
EPR experiments [M!ller & Brudvig (1991)]. It was identified as being located on
the D1 polypeptide by experiments using site-directed mutagenesis [Debus et al.
(1988); Debus et al. (1988b); Vermaas et al. (1988)]. a further tyrosine radical, Tyr
D is located on D2 and is also observable by EPR. It is not known where the Mn
cluster is located although some evidence suggests it is placed on the D2
polypeptide [Rutherford (1989)]. Other PSII polypeptides have not been extensively studied in this way as there are no known structures available for proteins
of sufficiently homologous sequence.
SMALL-ANGLE X-RAY SCATTERING A N D NEUTRON SCATTERING
STUDIES
Small-angle X-ray scattering [Kreutz, W. (1970); Sadler, et al. (1973)] and
neutron scattering [Worcester, D. L. (1975)] have been used to study photosynthetic membranes. These experimental techniques can give information about
the profile of density across stacked biological membranes. Using different
methods to interpret their data, Kreutz et al. (1970) and Sadler et al. (1973) found
the distribution of electron density of thylakoid membranes from small-angle
X-ray scattering with a stacked double membrane model with a repeat of about
160-170/k. In each case, information was obtained for the distribution of density
normal to the membrane plane. It was found that the width of a central region
between the stacked membranes could be varied by drying [Kreutz et al. (1970)]
or by changing experimental conditions in the centrifugation procedure used
[Sadler et al. (1973)] suggesting that this was representative of the lumenal space.
In neutron diffraction studies [Worcester, D. L. (1975)] of Euglena photosynthetic
membranes equilibrated with D20, a relatively intense peak was obtained at a
Bragg spacing of 85 A. These various studies are consistent with a structure
consisting of alternating hydrophilic and hydrophobic regions of about equal
thickness. The repeat spacing of the membrane pair was 170 ~. Since D20 readily
penetrates both the lumen of the thylakoids and between adjacent thylakoids (the
so called partition region--Fig. 1), it was concluded that the partition region was
not hydrophobic; as in this case it would be expected that the D20 layers would
be separated by membrane pairs rather than single membranes and the first
diffraction order of the 170 ]k repeat would be more intense than the second.
At the time at which most of these diffraction measurements were made,
little was known about the biochemical composition of the grana stacks compared
to the rest of the thylakoid membrane, but it is now known that the stacks are
highly enriched in PSII complexes which are laterally segregated from photosystem I and the other major membrane protein complexes, (the cytochrome b/f
complex and the chloroplast H+-ATPase). The repeat distance contains two
membranes with opposing surfaces, and essentially represents two thicknesses of
Photosystern II
165
PSII units, although clearly some interdigitation of PSII may occur, and hence
8 nm may be a slight underestimate of PSII thickness. The lipid composition of
the thylakoid membrane consists mainly of neutral lipids (di- or mono-galactosyl
diacyl glycerides) [Quinn & Williams, (1978)] and the small dip in density in the
neutron and X-ray diffraction measurements is likely to represent the centre of
the lipid bilayer. The predicted density profiles also showed a very strong
asymmetry with a far greater density of scattering matter on the lumenal side of
the membrane. Thus PSII is likely to have an unequal distribution of mass across
the thylakoid membrane, with a bias towards the lumenal side of the complex. It
has been known for at least 10 years that processes associated with photosynthetic
water oxidation are located on the lumenal side of PSII, and that extrinsic
polypeptides of mass 33, 23, 17 and 10kDa all have a lumenal location
[(Vermaas, (1993)]. In addition, the CP47 and CP43 subunits have large predicted
lumenal domains (>200 amino acids in size) but only small stromal domains. The
picutre that emerges from the biochemical and polypeptide analysis therefore
appears to agree with the earlier biophysical measurements.
ELECTRON MICROSCOPY
Ultrathin Sections
Ultrathin sections of thylakoid membranes examined by transmission EM
revealed their unusual morphology of interconnected membrane sheets (Fig. la).
Membranes are invariably associated in pairs, and these form a flattened sac
which encloses a space (the lumen). On the outside of the sac is a larger space,
the stroma which is bounded by the inner and outer chloroplast envelope
membranes. About half of the total thylakoid membrane area is found in
specialised regions where the flattened sacs closely stack on top of one another.
These grana are approximately circular in cross-section with a diameter of about
500 nm, and they can consist of up to about 20 stacked sacs. The unusual
morphology of these stacks makes them quite resistant to physical disintegration
by sonication or by French or Yeda pressure cell, and they also resist the
solubilisation action of some detergents. These characteristics have been exploited to produce relatively pure preparations of grana membranes.
Freeze-Fracture/Freeze-Etch Studies
Freeze-fracture and freeze-etch studies have also been employed to investigate thylakoid ultrastructure. At low temperatures, preparations of biological
membranes fracture preferentially along the hydrophobic phase of lipid bilayers,
revealing an exoplasmic and a protoplasmic leaflet. Membrane proteins remain
attached to the lipid layer in which they are more firmly anchored. Shadowing of
the faces reveals rounded particles with little or no substructure. This can be
improved by freeze-etching, during which ice on membrane surfaces adjacent to
fracture faces is etched away by sublimation. The intramembraneous particles
166
Nicholson,Ford, and Holzenburg
exposed by the freeze-fracture technique are generally accepted to correspond to
integral membrane proteins [Staehelin, L. A. (1975); Khodadad et al. (1986)].
However, it is possible that some particles observed by the method can also occur
due to artefacts created at various stages of the freeze-fracture procedure or may
arise from aggregates of lipids [Khodadad (1986)] as freeze-fracture particles have
been observed in liposomes lacking protein [Verkleij et al. (1979); Verkleij et al.
(1979b); Verkleij et al. (1980)]. Freeze-fracture studies of thylakoid membranes
initially only identified two distinct freeze-fracture faces [Henriques & Park
(1976); Arntzen et al. (1969)]. However, later work identified four different
freeze-fracture faces, distinguished by differences in the size and shape of the
particles embedded in the different fracture faces [Goodenough & Staehelin
(1971)]. The four distinct fracture faces are known as the PFu (protoplasmic face,
unstacked membranes), EFu (exoplasmic face, unstacked membranes), PFs
(protoplasmic face, stacked membranes) and the EFs (explosmic face, stacked
membranes) [Branton et al. (1975)]. Earlier terminology described them as the
Bu, Cu, Bs and Cs fracture faces respectively [Goodenough & Staehelin (1971)].
Protoplasmic refers to the stromal leaflet of the lipid bilayer whilst exoplasmic
refers to the lumenal leaflet (Fig. lb).
Biochemical analysis of stromal and granal membrane fractions suggests that
grana contain 80-90% of PSII [Andersson & Anderson (1980) as well as 70-90%
of LHCII [Kyle et al. (1983)] whereas 90% of PSI is found in the unstacked,
stromal lamellae. In early studies [Staehelin (1975); Staehelin et al. (1976);
Goodenough & Staehelin (1971); Branton & Park (1967); Miller & Staehelin
(1973); Ojakan & Satir (1974)] the freeze-fracture faces originating from the
stacked areas of the thylakoid membranes from spinach revealed two sizes of
particles; large particles of about 160 to 170 & cleaving with the EFs face and
small particles of about 80 ]k with the PFs and the PFu faces. It is now thought
that the larger EFs particles correspond to PSII complexes consisting of PSII core
together with associated light-harvesting proteins [Armond et al. (1977)]. In
studies of wild-type barley thylakoids the freeze-fracture faces had a similar
appearance to those observed for spinach membranes [Simpson (1978)]. In the
EFs fracture face a high density of the larger particles could be observed against a
smooth background. Smaller and less densely packed particles were observed in
the EFu face (which was continuous with the EFs face) and this face had an
undulating background containing numerous small pits. Several authors have
described two [Staehelin (1976)] or more [Armond et al. (1977)] maxima in size
distribution histograms of EFs particles. Differences in reported densities and
particle sizes may be due to different shadowing and counting methods [Kuhlbrandt ~1987)]. Closely packed particles appear to be partly obscured by unidirectic)na! shadowing; ~but revealed by rotary shadowing [Simpson (1979)]. The EFu
and PFu freeze-fracture faces, originating from unstacked, stromal membranes
are connected to the fracture faces of stacked, granal membranes, but differ in
particle size and density. The EFu face has the lowest density of particles and the
average size of EFu particles is about 110 ~. The complementary PFu shows
densely packed 105 ]k particles. Some authors distinguish two size classes of PFu
particles of 70-80 ~ and 105-120/~ diameter [Staehelin (1976); Armond et al.
Photosystem II
167
Freeze Etch
Freeze Fracture
Shadow
~\~,
Shadow
#
ESs Face
""
PFsFractureFace
i ( a ) Etch
Fracture-ill,,-~ ~ ~'~ t~"~-~ ~ ~
Plane
=
i
Fracture
Plane
(a)
Fracture
Plane
(b) Etch
I/'l -
~ , % ~ , - -~
~/~
Shadow ~ '
~,~
~,e1
EFs FractureFace
/~r162
y
PUC
PSs Face
Shadow S /
Replicafor EM
Fig. lb. Schematic view of freeze-fracture and freeze-etch electron microscopy methodology
showing the various fracture planes and surfaces discussed in the text for grana or stacked
thylakoid membranes. In freeze-fracture (left) frozen thylakoids are fractured along lines of
weakness, splitting the membrane down the middle. Two fracture faces are exposed, one where the
lumenal leaflet of the lipid bilayer remains behind (the EFs face), the other where the stromal
leaflet remains (PFs face). Protein particles are also present on the fracture faces, and different
populations of particles exist for each face. Current dogma implies that integral membrane protein
complexes separate into the face into which they are more deeply embedded, and for stacked
thylakoids the small particles observed on the PFs face are thought to be due to light-harvesting
complexes, whilst the larger particles on the EFs face are assumed to be PSII core. Assignments
have also been made for PSI and other complexes in the non-appressed lamellae or unstacked
thylakoids with EFu and PFu fracture faces. In freeze-etch methodology (right), fracturing along
planes in ice close to membrane surfaces is followed by a limited etching away of a thin layer of ice
by sublimation under vacuum to reveal the membrane surface topography. In both freeze-fracture
and freeze-etch cases, shadowing of the surface (usually at an angle of about 45~) with evaporated
metal (typically Pt/C) allows surface details to be revealed. The thin heavy metal surface (black
shape) is stabilised by a second evaporation of carbon at an angle perpendicular to the specimen,
and this forms a replica of the original surface. The replica is then separated from the specimen by
alkali digestion and examined in the electron microscope. The replica for just one of the surfaces
(the EFs face) is shown.
(1977)]. In studies of rotary shadowed PFu particles, Simpson [Simpson (1983)]
observed two distinct classes in particle size of 103 by 125 * and 67 by 83 ]k.
Similar observations have been made of thylakoid membrane surfaces using
freeze-etching. Freeze-etch faces from the lumenal surface of stacked thylakoids
168
Nicholson,Ford, and Holzenburg
(termed ESs) were found to be covered with large particles [Staehelin (1976);
Miller (1976); Simpson (1978)] which are similar with respect to their size,
distribution and number to the large EFs particles. It is thought that the ESs
particles represent the lumenal portion of the same large membrane protein
complex as the EFs particles (Fig. lb). Four domains can be resolved in ESs
particles giving them a 'tetrameric' appearance similar to rotary shadowed EFs
particles [Simpson (1979)]. Similar particles have not been observed on the
lumenal surface of unstacked thylakoids (ESu); but their presence may be
obscured by the rough relief of this surface type [Staehelin (1976); Simpson
(1978)]. The stromal surface of unstacked thylakoid membranes (PSu) has a
population of 130-180/~ particles against a background of 70-100 .~ particles
[Miller & Staehelin (1976)]. Stromal surfaces of stacked thylakoids are rarely
observed as the fracturing does not separate appressed membranes [Kuhlbrandt
(1987)]. However, granal membranes de-stacked near 0~ to minimise intermixing of membrane components showed smooth, almost featureless patches on the
stromal surface which may represent areas of membrane appression [Miller
(1981)]. Further evidence concerning the identity of the EFu and EFs particles
was provided by studies of photosynthetic membranes from mutants of higher
plant species. A tobacco mutant lacking PSII was observed to lack EFs particles
and 'tetrameric' ESs particles [Miller & Cushman (1979)]. Similar observations
have been made in freeze-fracture EM of a PSII-less mutant of the green alga
C h l a m y d o m o n a s reinhardtii [Wollmann, et al. (1981)]. There have been extensive
studies of thylakoid membrane preparations from barley mutants [Simpson
(1979); Bassi et al. (1985)]. The chlorophyll b-lacking barley mutant, chlorina-f2,
is deficient in LHCII and thylakoid membranes are observed to have substantial
reduction in the number of PFs freeze-fracture particles. Histograms of the size
distribution of particles suggested that EFs particles from the mutant [Simpson
(1979)] were about 12% smaller than those observed in wild-type barley thylakoid
membranes which implies that these particles are partly composed of lightharvesting polypeptides. The appearance of the EFu, PFu and PSu particles from
the chlorina-f2 mutant were essentially the same as those observed in wild-type
membranes. It is thought that the PFs particles represent LHCII and that the
change in size of the EFs particles is due to the loss of other Chl a / b binding
polypeptides.
Studies of PSII-enriched membranes and purified photosynthetic membrane
proteins reconstituted into liposomes provided additional evidence concerning the
identity of the freeze-fracture particles [Dunahay et al. (1984)]. Duhahay et al.
investigated four PSII-enriched thylakoid preparations using freeze-fracture EM.
The four different preparations could be divided into two different types on the
basis of the appearance of freeze-fractured membranes. PSII-enriched membranes produced by TX-100 treatment (i.e. the so-called BBY membranes-Berthold et al. (1981)) were found to have their lumenal or ESs surfaces, from
intact thylakoids, exposed. In contrast, membranes in PSII preparations from
digitonin treatment did not have this inside out orientation. Thylakoid membrane
preparations enriched in light-harvesting complex have been studied using
Photosystem I]
169
freeze-fracture EM, supporting the identification of the smaller (approximately
80 ~ diameter) PFs and PFu particles as the light-harvesting complex [Miller &
Cushman (1979); Lyon & Miller (1985)]. Soybean lecithin liposomes with
reconstituted LHCII (from pea and barley) were found to have particles of
similar size [McDonnel & Staehelin (1980)] on both fracture faces. Most of the
particles observed were arranged randomly, while some of the particles were
packed into hexagonal arrays. Further studies suggested that the LHCII particles
were also exposed on the surface of the membranes. Freeze-fracture of purified
barley LHCII in liposomes revealed particles similar in size and shape to PFs
particles [Simpson (1979)]. Photosynthetic membranes containing 2D arrays of
LHCII induced by Triton Xl14 were studied [Lyon & Miller (1985)]. The
membranes with crystallised LHCII were found to have hexagonal arrays of
LHCII which are observed on both the PF and EF faces with 12.5 nm spacing and
with individual particles of about 90 ~ in diameter.
PSII membranes given Tris-washing Or salt-washing treatment to remove the
various extrinsic polypeptides have been studied by using the freeze-etch
methodology [Seibert et al. (1987)]; Simpson & Andersson (1986)]. Washing
spinach PSII membranes with a buffer containing 0.25 M NaC1 removed the
16 kDa and some of the 23 kDa polypeptide, resulting in vesicles with a surface
containing essentially unchanged "tetrameric" ESs particles; but with a slightly
reduced surface relief [Simpson & Andersson (1986)]. After removal of the
16 kDa a n d the 23 kDa polypeptides with 1M NaC1 washing, ESs particles were
still present; but they had lost their "tetrameric" appearance. Washing with
1M CaC12 or alkaline 1M Tris resulted in particles which lost their "tetrameric"
appearance. CaC12 and Tris washing are thought to remove the 17, 23 and 33 kDa
polypeptides; Tris washing is thought to also remove most of a 10kDa
polypeptide [Ljunberg et al. (1984)]. Reconstituting the CaCI2 washed membranes
with a crude extract of the extrinsic polypeptides resulted in recovery of about
28% of the oxygen-evolving activity of the control membranes and in the
re-appearance of the four lobed ESs particles. In similar work, Seibert et al.
(1987) studied spinach PSII membranes after various washing procedures using
freeze-etch microscopy. In unwashed membranes ESs particles were observed
both in random orientations and in 2D arrays (with lattice dimensions of 17.5 nm
b y 20.4 nm, ~, = 90~ Treatment with 0.25 M NaCl in order to remove the 17 kDa
polypeptide resulted in ESs particles similar to those in control membranes but
with a slightly lower surface relief. Removal of the 17 and 23 kDa polypeptides
with 1M NaC1 resulted in further lowering of the height of the particles and ESs
particles with either "tetrameric" or "dimeric" appearance were observed.
Tris-washing to remove the 17, 23 and 33 kDa polypeptides resulted in a further
reduction in the height of particles and most of the ESs particles were found to
have a "dimeric" substructure. The Tris-washed, "dimeric" ESs particles were
estimated to be of height 6.1 nm and had similar diameter to particles in control
membranes. Particles in control membranes were of estimated height 8.2 nm.
Reconstitution of the washed membranes with the three extrinsic potypeptides
resulted in a recovery of up to 63% of the original oxygen-evolving activity and
Nicholson, Ford, and Holzenburg
170
the reappearance of multimeric ESs particles. Recently, a preparation of
thylakoid membranes containind 2D ordered arrays of PSII from the barley
mutant viridis-zb63 (lacking PSI) has been investigated by freeze-etch EM using
rotary and unidirectionally shadowed specimens together with image analysis
[Miller & Jacob (1991)]. The particles in the ordered arrays were approximately
16 nm by 22 nm in size. These particles are similar in size and appearance to the
EFs freeze-fracture and ESS freeze-etch particles, thought to represent PSII,
previously observed in wild-type thylakoid membranes not enriched for any
particular photosynthetic complex. Lattice parameters are not quoted in the
paper [Miller & Jacob (1991)] but they are approximately calculated from the
images presented to be 14 by 19 nm with 3/-- 81 ~ The particles, observed in both
reports, appear to have four major domains and were reported to have had
two-fold rotational symmetry. Further processing of images from the unidirectionally shadowed specimens was carried out in order to recover the surface
contour. [Miller & Jacob (1991); Smith & Kistler (1977)]. Close packed, small
particles were observed on the stromal PFs face as expected, and these were also
arranged in the same regular 2D lattice. The packing of the PFs particles suggests
that lattice contacts between PSII complexes are taking place on this side of the
membrane in the 2D-ordered arrays, especially along the a axis of the 2D crystals
where a large channel is observed between rows of the EFs particles.
Freeze-fracture studies of thylakoids from cyanobacteria have revealed that
particles of about 10nm size predominate in the exoplasmic fracture face
[Giddings et al. (1983); Golecki & Drews (1982); Lefort-Tran et al. (1973); Lichtle
& Thomas (1976); Morschel & Muhlethaler (1983); Neushul (1970)]. Particles of
about 10 nm size were observed in the fracture faces of liposomes containing
cyanobacterial PSII complexes [Morschel & Schatz (1987)]. Hence, it is likely that
the 10nm EF particles observed in cyanobacterial thylakoids represent
PSII complexes. Other evidence supports the same conclusion. A correlation
has been found between PSII activity and the number of 10nm EF particles
present both in wild-type and a phycobilisome-deficient mutant of the red
alga Cyanidium caldarurn [Wollman (1979)]. Heterocyst thylakoids do not have
PSII activity and also lack 10nm EF particles [Giddings & Staehelin (1979)].
Studies of the 10nm EF particles of cyanobacterial thylakoids suggested that
they were arranged in dimers of 10nm by 20nm. The dimers were attached
to one another at their longitudinal faces to form rows of variable length with
centre-to-centre spacings for the particles of about 11nm. The large spacing
observed between neighbouring rows (45 to 75 nm) was interpreted as being
due to parallel phycobilisome rows at the outer surface of the thylakoids
which are known to have a similar centre-to-centre distance [Morschel & Schatz
(1987)].
Negatively Stained PSII
Electron microscopy of negatively stained specimens has proved to be a
powerful tool for the structural elucidation of PSII. In negatively stained
specimens, contrast is imparted by solutions of heavy metal salts which upon
Photosystem II
171
drying form a glassy cast around the protein. It is assumed that the metal salt
solution is able to occupy the hydrated regions in and around proteins, i.e. protein
corresponds to stain-excluding areas. Electron microscopical studies of negativelystained PSII-enriched membranes, detergent-solubilised PSII complex, PSII core
complex and also of CP47, D1, D2 cyt b559 sub-complex have all been reported.
A major omission from this list (as far as we are aware) is the reaction centre
D1/D2/b559 preparation. We are not sure why there are no reports of EM of this
material, but this review may prompt the study, or perhaps prompt someone to
inform us of the existence of such studies. The maximum expected resolution for
negatively stained specimens is usually around 2nm, although under optimal
conditions this may be extended to about 1 nm [(Unwin & Klug (1974)].
Spinach PSII core complexes with high oxygen-evolving activity were purified
and studied by EM [Haag, et al. (1990)]. Etectrophoretic analysis suggested that
the purified PSII core complexes were composed of the 47 kDa, 43 kDa, D1 and
D2 polypeptides and the two subunits of cytochrome b559 and that there was
some contamination with the chlorophyll binding polypeptide CP29. Image
analysis by single particle averaging [Boekema et al. (1986)] was used to obtain
improved images of side-on and face-on views of the complexes (distinguished by
higher and lower contrast particles respectively). The side-on views had an
ellipsoid appearance with approximate dimensions 15.6 nm by 7.3 nm and had a
small protrusion estimated to be between 1.5 nm and 3.3 nm in height which was
ascribed to the 33kDa extrinsic polypeptide. Particles interpreted both as
monomeric and dimeric PSII were observed in the preparation. The face-on
projection of the monomer had a triangular shape and was approximately 15.6 by
10.6 nm (see Fig. 2(k)). 54 particles were used to obtain the top view average and
30 particles to obtain the final side view average. The authors do not quote any
resolution estimate. It has been suggested [Dekker, et al. (1990)] that the
triangular particles were due to contamination by LHCII, however, the particles
appear to be the wrong size to be consistent with this interpretation; as the
diameter of the LHCII trimer is reported to be 7.3 nm in high-resolution studies
[Kuhlbrandt & Wang (1991)]. More recent work by the same group has focused
on the larger particles in the PSII core complex preparation [Haag, et al. (1990)]
which were postulated to be dimeric [Boekema et al. (1994)]. The authors
maintain that the PSII core complex is usually present in vivo in the dimeric form
(i.e. it has two complete sets of PSII polypeptides, pigments and electron transfer
components) and the whole complex is predicted to have two-fold rotational
symmetry. The observed particles of dimension 17 by 10 nm exhibit four distinct
stain-excluding areas (Fig. 2(b) a n d ( h ) ) . The resolution was estimated to be
about 2 nm using an unstated criterion. The authors also characterised a larger
particle from spinach (Fig. 2(a)) of approximately 30 by 16 nm (face-on), a height
of 8.8-9.3 nm in the centre and 6 nm at the outer ends (side-on) [(Boekema et al.
(1995)]. It was inferred that the central area of stain-excluding density could be
identified as the smaller PSII core dimer. The peripheral areas of density in the
image of the averaged large particle were attributed to LHCII and other
light-harvesting polypeptides. Various sub-populations of the particles in the PSII
core complex preparation have been identified [Haag et al. (1990)]. The
172
Nicholson, Ford, and Holzenburg
b
c
d
e
88
20 nm
Fig. 2. Schematic representations of various reported PSII
structures drawn to the same scale: (a) Supercore "dimer"
from spinach (Boekema et al., 1995). (b) Spinach core "dimer"
(Boekema et al., 1994). (c) Spinach core monomer in thin 3D
crystals (Dekker et al., 1990). (d) and (e) Cyanobacterial
monomer (d) and "dimer" (e) (Roegner et al., 1987). (f)
Tetrameric ESs particles (freeze-etch method) (e.g. Seibert et
al., 1987). (g) Spinach "dimer" in 2D arrays (Lyon et al.,
1983). (h) Cyanobacterial "dimer" (Boekema et al., 1994). (i)
Spinach monomer in 2D arrays (Holzenburg et al., 1992, 1993).
(j) Maize "dimer" in 2D arrays (Santini et al., 1994). (j)
Spinach monomer core (Haag et al., 1990). In the case of
structures derived from ordered 2D arrays of PSII in membranes (Lg, i,j), the unit cell dimensions are shown as a box
enclosing the PSII structure. All are derived from negativelystained specimens except (f). Two-fold rotational symmetry
was imposed for all the structures noted as "dimer". For
detergent solubilised PSII structures, the drawings depict the
complex in "face-on" view which was taken as being an
orientation of the complex where the plane of the paper
represents the original membrane plane.
h e t e r o g e n e i t y of the preparation m a y be due to deficiences in the purification
(which electrophoresis suggests is unlikely) or alternatively due to variable
aggregation states of PSII.
T h i n 3D crystals of the CP47, D1, D2, cyt b559 core c o m p l e x of spinach PSII
h a v e b e e n reported [ D e k k e r e t al. (1990)]. T h e crystals w e r e studied by E M and
w e r e suggested to have pgg s y m m e t r y in projection with a rectangular unit cell of
dimensions 23.5 nm by 16.0 nm. M o n o m e r s were reported r a t h e r than dimers and
the a u t h o r s suggest that a PSII p o l y p e p t i d e other than those present in the
purified sub-complex is required for the formation of dimeric PSII. T h e PSII
m o n o m e r s observed had an asymmetrical shape with dimensions of 10 n m by
7.5 n m with a height of 6 n m (Fig. 2(c)). T h e height was estimated f r o m side-on
views of the crystals. Specimen purity was assessed by gel p e r m e a t i o n c h r o m a t o g r a p h y which suggested that the p r e p a r a t i o n consisted of a h o m o g e n e o u s
PhotosystemII
173
population of particles. However, the crystals show a close similarity to thin 3D
crystals of LHCII reported by Kuhlbrandt (1983). Although the 2D space group
of the crystals was identified as pgg, they appear to be organised in a hexagonal
pattern with dimensions comparable to the hexagonal packing of LHCII. Hence,
it is possible that the space group was incorrectly chosen due to distortion of the
hexagonal lattice caused by the superposition of two or more crystalline layers
which can slide against each other. The micrographs shown do indeed suggest
such an arrangement, with multiple layers apparent by the different degrees of
electron scattering for different thicknesses of the specimen.
Ordered 2D arrays of spinach PSII core complex after apparent solubilisation
and reconstitution into double layered flattened vesicles were reported recently
[Lyon, et aL (1993)]. A 2D projection map of the negatively-stained crystals was
obtained using correlation averaging [Frank et al. (1981)] due to the inherent
disorder of the 2D arrays and a dimeric structure was assigned (Fig. 2(g)). The
resolution was estimated by the spectral signal-to-noise ratio method [Unser et al.
(1987)] to be about 1.7nm. Gel etectrophoresis showed that the crystal
preparation consisted of core polypeptides (i.e. the 47 kDa, 43 kDa, cytochrome
b559, D1 and D2 polypeptides) and lacked the peripheral OEC polypeptides. The
gel electrophoresis also suggested contamination of the crystal preparation with
LHCII. The PSII monomers observed in the projection map were of approximate
dimensions 9.7 nm by 5.3 nm. The 2D crystals had unit cell dimension 11.5 by
16.1nm with y = 7 5 . 3 ~ Assuming a density of 770Da/nm 3 [Unwin & Ennis
(1984)] and crystal thickness of 9.8 nm the authors estimated the molecular mass
consistent w i t h the volume of the dimer to be about 810 kDa. A ro~ationaI power
spectrum [Crowther & Amos (1971)] was computed and found to be consistent
with the crystals having 2-fold symmetry. It was concluded that the space group of
the 2D crystals was p2. Only four distinct areas of density could be discerned per
monomer which is surprising when considering the resolution reported by the
authors. Another unusual feature of this work was the apparent removal of
LHCII from the preparation with concentrations of detergent, Triton X-100,
which would normally not solubilise the spinach PSII grana under the conditions
reported (2% w/v was used followed by 1% w/v). These conditions are, however,
reported to allow the isolation of PSII-enriched grana from barley thylakoids
[Simpson & Andersson, (1986)].
Electron microscopical studies have been made of negatively-stained 2D
crystals in PSII-enriched membranes from Z e a m a y s [Bassi et al. (1989); Santini et
al. (1994)]. The crystals were produced by stacking of maize PSII-enriched
membranes at pH 7.5 followed by further treatment with Triton X-100. The
authors presented a low resolution projection [(Bassi et al. (1989)] and later 3D
structure [(Santini et al. (1994)] of PSII with and without treatment of PSII
membranes by Tris-washing which removed the OEC polypeptides. Non washed
PSII was observed to have a four-lobed structure and two-fold rotational
symmetry was imposed in the map. An apparent square pattern with a repeat of
52 nm could be observed in electron micrographs of the 2D crystals. Numerical
diffraction patterns suggested that there were two superimposed rectangular
174
Nicholson, Ford, and Holzenburg
lattices rotated by 90~ with respect to each other in the membranes with lattice
dimensions 26nm by 18nm (3' =90 ~ (Fig. 2(j)). Layer-specific data were
extracted by Fourier filtering and the images of each layer were improved by
correlation averaging [Saxton et al., (1984)]. The method of Cjeka et aL [Cjeka et
al. (1986)] was used to identify the layer to which each of the averages belonged.
Averages obtained from the different layers were found to have spatial
resolutions of 2.2 nm and 2.4 nm (for the lower layer) using t h e Fourier ring
correlation criterion [van Heel (1987)] which is reported to provide a more
optimistic estimate of resolution than other methods [Unser et al. (1987)].
Averaged images of projections of the 2D crystals presented in earlier work by
the same group [Bassi, et al. (1989)] were estimated to have a resolution of 5.0 nm
as judged from the appearance of computed diffraction patterns. Prior to
Tris-washing, four main domains could be distinguished in the structure; whereas
the Tris-washed crystals revealed only two domains. The threshold used to
calculate the volume of the 3D reconstruction did not allow any lattice contacts to
be observed which appears to indicate either a low resolution or that too high
a threshold was chosen. The volume of the PSII complex with and without the O E C polypeptides was estimated as 640nm 3 and 600nm 3 respectively using estimates for the thickness of the complex, before (13.5nm)
and after Tris-washing (10nm). These values are probably an over estimate
as small-angle X-ray and neutron scattering studies suggest a thickness of
about 8.5 nm [Sadler, et al. (1973); Kreutz (1970); Worcester (1976)]. The mass
calculated was 630 kDa and 672 kDa for the Tris washed and untreated PSII
respectively.
Two-dimensional crystals of oxygen-evolving and Tris-washed spinach PSII
(in grana membranes) have been studied using image analysis to yield projection
maps [Holzenburg et aL, (1992); Holzenburg et al., (1994)] and 3D structures
[Holzenburg, A. et aL (1993); Ford et al., (1995)] at 3 nm to 1.8 nm resolution.
This work, which is discussed in more detail later, concluded that a monomeric
form of PSII exists in the native thylakoid membranes (Fig. 2(i)).
PSII from cyanobacteria has also been studied by EM [Rogner et al. (1987);
Dekker et aL (1988)]. The authors obtained images of negatively stained PSII
core complex (solubilised by dodecyl maltoside) purified from Synechococcus
elongatus. Although it is generally thought that native PSII exists in cyanobacteria
as dimers [Manadori & Melis (1985)] the EM studies of detergent solubilised PSII
revealed both dimers and monomers [Roegner et aI. (1987)] (Fig. 2(d) and (e))
and both forms were found to be active in oxygen evolution. The PSII dimers
were estimated to have dimensions of 1574 12.8nm (by subtraction of a
postulated detergent shell of about I nm thickness) and a height of 5.6 nm
(estimated from the average of the height of individual projections) and in
addition there was a 0.8 nm protuberance on the lumenal side of the membrane
which was assigned as the 33 kDa OEC subunit. In a face-on projection, the
monomer was found to be roughly oval in shape with dimensions of 12.3 by
7.5 nm after adjustment for detergent as before.
In conclusion, the different EM studies of plant PSII are apparently
inconsistent (see Fig. 2 for a gallery of PSII structures). For example the
PhotosystemII
17~
"supercore" complex (Fig. 2(a)) reported by Boekema et al. (1995) is much larger
than the membrane-native PSII structures (Fig. 2(0 (i) (j)), although it should be
smaller as it has lost part of the light-harvesting antennae proteins. The
cyanobacterial core monomer (Fig. 2(d)) reported by Roegner et al. (1987) is as
large as cyanobacterial and spinach dimers (Fig. 2(h) and (b) which were reported
later by the same authors [(Boekema et aL (1994)]. Similarly, the large triangular
core monomer (Fig. 2(k)) from spinach [(Haag et aL (1990)] is difficult to
reconcile with any of the other PSII structures. The most consistent structures are
for membrane-embedded PSII where a 4-lobed complex of about 20 nm by 17 nm
is reported (Fig. 2(f), (i) & (j)). The inconsistency of the detergent-solubilised
PSII preparations may be due to heterogeneous or contaminated PSII preparations or difficulties in interpreting the EM data correctly. However in our opinion,
the major factor governing this inconsistency is the effect of detergent-induced
oligomerisation and association which can occur after solubilisation of the native
membrane. Oligomerisation of PSII after solubilisation in detergent appears to be
common to both pro- and eukaryotic PSII. It is therefore important to study the
complex in its native membrane whenever possible, or if detergent-solubilised and
purified PSII is needed (as for cyanobacteria) then the oligomeric form should be
rigorously characterised.
THREE-DIMENSIONAL ARCHITECTURE OF PSII
Three-dimensional information can be obtained from 2D crystals by collecting EM data with the specimen tilted at various angles with respect to the
electron beam [(Amos et al., 1980)]. For a crystal which is only one molecule
thick the resulting Fourier transform along z* (the direction in reciprocal space
perpendicular to the 2D crystal plane) will be continuous. A plot of amplitude
and phase data for every measured lattice line (indices h, k) versus tilt angle will
give an approximation of the continuous transform along z* which can then be
sampled at regular intervals to yield the 3rd index (1) and thus structure factors in
three dimensions (h, k, 1) in the reciprocal lattice. The degree of scatter of the
data gives an indication of the accuracy of the data and provides an independent
estimate of the resolution cut-off. In Fig. 3, data for PSII 2D arrays is plotted in
such a manner for lattice lines (3, 0) and (2, - 8 ) (data from Ford et al., (1995)).
The spread of the data is clearly greater for the amplitude information than for
the phases, and also the scatter is generally greater for the weaker, higherresolution reflections. It is important to consider the effects of this on the
computed 3D map. Firstly, because the phase information is more reliable than
the amplitude information, the positions of domains in the 3D map are more
accurate than their volume. Since the 3D map is represented by contour lines or
surfaces at a selected density threshold, then we can replace the term "volume"
for 'denisty' in the previous sentence. It is therefore normal to display the 3D
map at different density thresholds in order to allow for the variability in the
density of different domains without the potential loss of information. Secondly,
176
Nicholson, Ford, and Holzenburg
o
18~
~
3,0
~ ~ 1 7 6
~
/
~ ...................
\::-o
_la0,I
I
.....,......
x ~ _ o
~ o/,,.
".......I
I
151
c=
<>
o
ampl.
o ='~ ~
o
o
~
o
o o
-0.05
0
z*
0.05
[,~l -~
2,-8
I
phase 0 I
.la0~
33
o
ampl.
0
-0.05
o
o
b.05
0
z*
(~1-1
Fig. 3. Variation and scatter of amplitude and phase data for two
selected lattice lines (h = 3, k = 0) and (h = 2, k = - g ) from a 3D data
set (Ford et al., 1995). Scatter for amplitude data is always larger than
for phases, and scatter for both phase and amplitude increases with the "
weaker reflections at the limit of resolution (see text). The z* axis
represents distance in reciprocal.space perpendicular to the 2D crystal
plane with units in reciprocal A, and the magnitude of z* values is
determined by the tilt angle as well as the h and k values of the
reflection. Values for the amplitude of the reflections are relative.
Smooth curves are fitted to the data and structure factors extracted at
defined points along z.
b e c a u s e t h e d a t a is n o i s i e r at h i g h e r r e s o l u t i o n , small a n d w e a k l y d e f i n e d d o m a i n s
must be treated with scepticism, and should preferably only be included in the
i n t e r p r e t a t i o n o f a n y 3 D m a p w h e n t h e y a r e c o n s i s t e n t l y f o u n d in t h e s a m e
p o s i t i o n i n c o m p l e t e l y i n d e p e n d e n t d a t a sets.
Photosyslem II
177
Discrimination between noise and structure in the 3D map is still a relatively
subjective process, and one should adopt the same criteria as X-ray crystallographers where density thresholds used for display are defined in terms of the
r.m.s, density in the entire unit cell. Thus thresholds might typically be used at
densities corresponding to 1.0 or 1.5 times the standard deviation above the mean
density level, where the mean density is thought to approximate to the density
due to the solvent (water is the major component in most 3D crystals of proteins).
For negatively stained specimens, these thresholds can still be used, but in this
case the solvent is essentially replaced by the heavy metals comprising the
staining solution, and often positive staining occurs for some regions of the unit
cell. This means that the r.m.s, values of unstained and stained maps are not
directly comparable because positive staining will give rise to apparent strong
negative domains in the computed map (not observed for unstained crystals), and
this means that the r.m.s, values are larger than would perhaps be anticipated
(because real structure due to protein can be both positive and negative). These
problems can make the interpretation of maps from negatively stained specimens
difficult. The representation of the PSII 3D map shown in Fig. 4 therefore uses
three different density thresholds for the display at 0.5, 1.0 and 2.0 times the
standard deviation above the mean density. At the lowest threshold (blue netting)
the small domains that surround the circumference of the main body of the
complex must be considered to be at the noise/structure borderline and should be
treated with suspicion. However some of them are consistently found in
independent 3D maps and therefore meet the criterion stated above.
There are only 3 reports of 3D structures of PSI1 [Holzenburg et al. (1993);
Santini et aL (1994); Ford et al. (1995)]. A comparison of the 3D maps of
Holzenburg et al. (1993) and Santini et aL (1994) reveals consistent features
between the two, although the map of Holzenburg et al, is clearly much more
detailed (see Fig. 2). The most obvious common features are the four lumenal
domains of the complex as well as the asymmetric distribution of mass with
respect to the membrane. The four lumenal domains are given the Roman
numerals I-IV by Holzenburg et aL (1993) in order to define them. In the model
of Santini et a t , the 4 domains, each about 4 nm in diameter, are the finest details
resolved, and on the stromal side of the complex, only two 6rim diameter
domains are shown and lattice contacts cannot be discerned. Tris washing to
remove extrinsic snbunits was also reported by Santini et aL, although this led to
an apparently inverted structure consisting of two featureless 6 nm diameter
domains on the lumenal side extending into four smaller domains on the stromal
surface. It is not obvious why the maps presented by Santini et al. (1994) contain
so little detail. The resolution reported by the authors (<2 rim) ought to be
sufficient to resolve subunits larger than about 15 kDa, of which there are at least
20 in PSII). However, it is possible that by imposing 2-fold symmetry in their data
analysis, fine detail could be smeared out. This would hold true if the space group
is not p2 but pl, as contended by Holzenburg et al. (1993). In order to discuss
PSII architecture in finer detail, we must now consider the 3D maps of
Holzenburg et al. (1993) and Ford et aL (1995).
178
Nicholson, Ford, and Holzenburg
a
b
C
,4
Fig. 4. 3D structure of native PSII depicted with: (a-c) sections from the 3D map using contour lines
to show density and (d-e) with chicken-wire presentation using red, yellow and blue colours to
represent density. Section (a) is taken from the middle of the lumenal side of the complex and shows
the characteristic four-domain appearance and central cavity, Section (b) is taken from a position
close to the lumenal membrane surface and shows the merging of the four domains, loss of the cavity
and appearance of small peripheral domains (arrows). Section (c) is laken from the slroma] side of the
Photosystem II
179
Beginning with the lumenal side of native PSII (Fig. 4a, d & e), the 3D data
shows that the four domains (I to IV) do not project directly upwards like
columns, but rather lean inwards towards the centre of the complex. Thus the
four domains surround a cavity which has a narrow opening at the top, but which
opens out into an egg-shaped chamber that is about 5 nm in length, with a height
of about 3.5 nm. If the observer moves from the lumenal face towards the
membrane surface, the four domains begin to merge, with domains IV and II
shifting towards a more central location whilst domains I and III fade out (Fig.
4b). At this point, a new feature in the 3D map becomes apparent around the
periphery of the complex (Fig. 4b, arrows). We can now observe the partial
resolution of 8 protruding domains that are equidistantly spaced (rather like
stubby spokes sticking out from the hub of a wheel). At this level in the map,
contacts along the b axis (the longer one) of the lattice can be discerned, and they
appear to involve the interaction of these spoke-like domains. Lattice contacts
along the a axis are still not apparent, however. As the observer moves through
the membrane and over to the stromal side of the complex, the map becomes
very detailed (Fig. 4c). At the centre of the complex lie two domains (A and B)
that derive from lumenal domains IV and II respectively. Domain A is larger with
a diameter of about 6 nm. Surrounding domains A and B are many discrete small
domains, and 8 of them derive directly from the 8 projecting spokes observed on
the lumenal side of the complex (they are better resolved on the stromal side, Fig.
4c). Contacts along the a axis (the shorter one) can now be seen, and again,
domains connected to some of the spokes are involved in these contacts. There is
no cavity on the stromal surface, which in comparison to the lumenal surface of
the complex, is quite smooth and flat, and extends only about 1 to 2 nm out from
the predicted edge of the lipid membrane (see Fig. 6). It is clear from the EM that
the lumenal surface is in contact with the specimen support film. This means that
the stain is much deeper on the lumenal side of the complex, leading to higher
apparent densities for the lumenal domains. This presents problems for the
display of the 3D map because display threshold for the lumenal face should be
different to that used for the stromal face, which is impractical. Due to this, the
stromal side of the complex appears truncated at the highest (red) thresholding
which is really only of use for the lumenal side. Although the stromal face in the
map has lower densities on average, previous experience suggests that this face is
less likely to suffer from distortions due to flattening because it is not in contact
Fig. 4. Continued--complexand shows the two domains A and B as well as many smaller peripheral
domains (arrowheads). In all cases the complex is viewed from the stromal side. For scale, the unit cell
dimensions are 16.8 by 18.9nm. In panels d and e the complex is represented by chicken-wire which
correspond to densities at 0.5 (blue), 1.0 (yellow) and 2.0 (red) times the standard deviation above the
mean density level. In panel (d), the complex is viewed from the stromal side, face-on and in (e) a side
view is shown (the lumenal face is up in the image). The side-view reveals the intramolecular lumenal
location of the cavity and also shows protein density fading out and then reappearing on the stromal
side of the complex (double arrows), suggesting that this is the hydrophobic portion of the complex
embedded in the lipid bilayer (see Fig. 6). Scale bar = 5 nm (panels d and e).
180
Nicholson, Ford, and Holzenburg
with the support film, and therefore the potential resolution for this face may be
at least as high as the other face.
Recent data on Tris-washed PSII (Ford et al. 1995) has also resulted in a 3D
map which is discussed here. The resolution reported for the Tris-washed map
was 1.8 nm compared to 3 nm for the native structure. These values refer to
resolution in the plane of the crystal. Resolution perpendicular to the crystal
plane (along z) is worse because of limitations that are due to several problems
concerned with collecting data with the specimen at high tilt angles relative to the
electron beam. Resolution along z for the Tris-washed and native forms are
estimated as about 3 nm and 5 nm respectively.
Figure 5 shows a 3D representation of the structure using sections from the
map (Fig. 5a & b), red and blue netting to represent different density thresholds
as before (Fig. 5c), and also a surface-rendering (Fig. 5d). Tris-washing was found
to affect the lumenal side of PSII much more than the stromal side, and in Fig. 5a,
c and d it can be readily observed that the cavity of the native lumenal structure
has completely disappeared after Tris-washing. Domain IV appears unaffected by
the treatment, whilst domains I and III are lost. Domain II is only partly lost, but
what remains appears to be in a shifted position that would place it towards the
bridging region between domains III and II in the native structure. In the
Tris-washed map, this domain was therefore named V, (with domain II in the
native structure being predicted to be formed partly from domain V plus some
other protein component that is removed by Tris-washing). In the Tris-washed 3D
structure, data was obtained for crystals which were attached to the support film
by their lumenal faces (as before), but data was also obtained for stromallyattached crystals. The distribution of density is therefore much more even in the
map (compared to the native map), making display easier. A 3D difference map
(Fig. 6) shows quite clearly that there are three major lumenal domains that are
removed by the Tris-washing and these evidently help to form the egg-shaped
cavity in the native structure. Since Tris-washing is known to remove the three
Fig. 5. 3D structure of Tris-washed PSII depicted using contoured sections from the 3D map
(a-b), netting at 0.6 (blue) and 2.4 (red) times standard deviation above the mean density (c)
and finally a non-transparent surface relief at 1.6 times the standard deviation above the mean
(d). In panels a and b the viewing direction is from the stromal side of the membrane and the
unit celt dimensions are 17.7 by 20.1 nm. The lumenal section (a) shows two major domains (IV
and V) with the loss of the central cavity. The stromal section (b) shows the emergence of a
domain (VI) that lies on a local pseudo-twofold rotation axis that relates domains IV and V.
Peripheral domains that coincide in the native and Tris-washed maps are marked by
arrowheads. In panel (c), domains IV-VI are highlighted, viewed, (as in a & b) from the
stromal side. There is a discernible core structure which is mainly enclosed by the red netting.
This is surrounded by many smaller peripheral densities that are weaker and are enclosed by
the blue netting. As in the native structure, these peripheral domains are involved in making
the lattice contacts. In panel (d) the observed looks down onto the surface of the complex from
the lumenal side. The approximate positions of missing domains I, II and III are marked by
arrows. Note that domain III would be expected to be located underneath a thin flap of protein
density that bridges between domains IV and V in the outer lumenal regions of the complex.
Scale bars represent 5 nm in panels c and d.
Photosystem II
a
....
9Y
181
b
4~
.....
i
182
Nicholson, Ford, and Holzenburg
Fig. 6. Fourier vector difference map calculated from native PSII minus Tris-washed PSII (blue
netting at 1.9 times the standard deviation above the mean density) superimposed on the native PSII
3D structure (red netting at 1.0 times the standard deviation above the mean density). Three domains
(representing the positions of domains 1--far side of cavity, If--near side & III--near side) are
evident in the difference map on the lumenal side of the complex and they surround the central cavity
of the native structure. Also shown (white circles) are the expected boundaries of the lipid bilayer (see
text). The lumenal surface is uppermost in the figure. Scale bar = 5 nm.
O E C subunits of mass 33, 23 and 17 k D a , it was concluded that the three peaks in
the difference m a p represent the locations of these extrinsic subunits. Furtherm o r e it was postulated that the intramolecular cavity in the complex could house
a special microenvironment, perhaps with elevated Ca 2+ and C1- levels, which
allowed optimal oxygen-evolution activity in PSII [Holzenburg et al. (1994); Ford
et al. (1995)]. R e m o v a l of the O E C subunits would therefore lead to a loss in the
activity of the complex that could be regained by adding exogenous CaC12.
On the stromal face in the Tris-washed PSII structure, several small discrete
domains around the periphery of the complex are again observed (Fig. 5b), and
the positions of several of these domains coincide with those found in the native
PSII structure (arrowheads). It has b e e n postulated that these peripheral domains
could represent L H C I I and associated light-harvesting antennae proteins (Holzenburg et al. 1993). Although these w e a k e r domains lie at the structure/noise
boundary in the map, at least 6 of t h e m a p p e a r to be at equivalent positions in the
native and Tris-washed 3D maps, thus strengthening their localisation. A further
central domain appears which is flanked by domains IV and V. This domain has
been termed domain VI and lies on a pseudo-2-fold axis that relates domains I V
and V by a 180 ~ rotation. In the lower resolution native structure [Holzenburg et
al. (1992)], domains IV and V I were not resolved and appeared as a single
PhotosystemII
183
domain (domain A, Fig. 4c). As stated earlier, a pseudo-symmetry is expected for
the PSII core structure since there is considerable sequence homology between
D1 and D2 as well as between CP47 and CP43.
SIZE A N D OLIGOMERIC FORM OF PSII
Perhaps surprisingly, there is still considerable debate over the oligomeric
form of plant PSII in the native membrane (see Fig. 2), and most of the
arguments for and against a dimeric structure have already been rehearsed in this
review. In our view, the most telling evidence against the dimeric model comes
from considerations of the predicted size of the monomeric complex versus the
available space in the unit cell of the 2D crystals observed in grana membranes.
By size, we do not only refer to the molecular mass, but also the size of the
photosynthetic unit (i.e. the number of chlorophylls associated with each PSII
electron transfer chain). Firstly, for the molecular mass calculations, the lowest
threshold used in 3D data analysis for the complex gives a volume of about
1000 nm 3 (a lower volume of 800 nm 3 was calculated for the Santini et al. 3D map)
and taking standard values for partial specific volume, a mass that could be
enclosed by this volume of about 800 kDa is calculated (Santini et al. calculated
700kDa). A dimeric PSII complex would have a mass of at least 1500kDa
(Holzenburg et al., 1993). Secondly, we consider photosynthetic unit size. As
stated earlier, the grana-located PSII complexes have at least 8 and at most 20
peripheral light-harvesting polypeptides (mainly LHCII) associated with each
photosystem II monomer. Thus a dimeric PSII complex would have at least 16
peripheral LHC polypeptides which have to be occupied in the PSI! unit cell with
an available area in the membrane of 17 • 19 nm (323 nm2). Since the membrane
area occupied by a single LHCII monomer is known from high resolution
electron crystallography to be about 24nm 2 (Kuhlbrandt et al., 1994), we can
calculate the maximum number that could possibly be packed into the PSII unit
cell. The answer is about 14, and thus, even ignoring the requirement for PSII
core complex and lipid, there is not enough room in the unit cell to house even
the peripheral light-harvesting proteins of a d i m e r i c complex. The native form of
PSII must therefore be concluded to be monomeric, with what must necessarily
be a very tightly packed structure in the membrane crystals. The core complex
must make a relatively compact footprint of about 130 nm 2 or less in the lipid
membrane.
SPECULATIONS A B O U T SUBUNIT ARCHITECTURE
The two largest core polypeptides, CP47 and CP43, are expected to have
extensive lumenal domains which are not removed by Tris-washing. It therefore
seems a reasonable guess that domains IV and V are probably the lumenal
184
Nicholson, Ford, and Holzenburg
portions of these two polypeptides although which is which beyond speculation
(at present). The core complex consists of CP47, CP43 and the reaction centre of
D1, D2 and cytochrome b559. There is a discernible core in the structures
[(Holzenburg et al. (1993); Ford et al. (1995)]. It has therefore been postulated
that domain VI is the location of the reaction centre, which sits roughly on the
2-fold pseudosymmetry axis and is flanked by the putative CP47 and CP43
domains. Around the core density in the maps of Holzenburg et al. (1993) and
Ford et al. (1994) are peripheral densities that (by their size, number and
location) may be light-harvesting polypeptides such as LHCII. From calculations
such as those described above, we expect at least 8 light-harvesting proteins to lie
in a shell that is involved in the lattice contacts. Some of the peripheral domains
in the maps shown are not intimately associated with the main body of the
complex, and although these weaker domains could be noise in the maps, it could
be possible that these may be a mobile portion of LHCH that can be reversibly
phosphorylated and migrate to and from the non-appressed thylakoid
membranes.
FURTHER DEVELOPMENTS
Future improvements in knowledge of the structure of PSII will depend on a
close collaboration between biochemists and structural and molecular biologists.
The best hope for very high (<3 ]k) resolution data probably lies in X-ray
crystallography of higher plant or cyanobacterial core complexes, but this is
presently limited by the quality and stability of the crystals. Electron microscopy
can provide a much more rapid route to obtain structural data at low to high
resolution (50 to 3 ]k), and is also likely to be the only way of obtaining
information on the overall architecture of the intact complex in the membrane.
ACKNOWLEDGEMENTS
We wish to acknowledge Drs Svetla Stoilova, Ashraf Kitmitto, Mark
Rosenberg, Fiona Shepherd and Messrs Toby Flint and Richard Collins for vital
discussions concerning this work. The authors are grateful to the BBSRC, the
Royal Society, the Nuffield Foundation, the Akademie der Wissenschaften in
Goettingen, the Academic Development fund of the University of Leeds and the
Leeds Centre for Molecular Recognition in Biological Sciences (University of
Leeds) for financial support for their studies on PSII structure.
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