Photosynthesis Research 63: 195–208, 2000.
© 2000 Kluwer Academic Publishers. Printed in the Netherlands.
195
Minireview
Primary charge separation in Photosystem II
Jan P. Dekker∗ and Rienk Van Grondelle
Faculty of Sciences, Division of Physics and Astronomy, Vrije Universiteit, De Boelelaan 1081, 1081 HV
Amsterdam, The Netherlands; ∗ Author for correspondence (e-mail: dekker@nat.vu.nl)
Received 3 November 1999; accepted in revised form 10 February 2000
Key words: charge separation, disorder, exciton interaction, Photosystem II, reaction center
Abstract
In this Minireview, we discuss a number of issues on the primary photosynthetic reactions of the green plant
Photosystem II. We discuss the origin of the 683 and 679 nm absorption bands of the PS II RC complex and suggest
that these forms may reflect the single-site spectrum with dominant contributions from the zero-phonon line and
a pronounced ∼80 cm−1 phonon side band, respectively. The couplings between the six central RC chlorins are
probably very similar and, therefore, a ‘multimer’ model arises in which there is no ‘special pair’ and in which
for each realization of the disorder the excitation may be dynamically localized on basically any combination of
neighbouring chlorins. The key features of our model for the primary reactions in PS II include ultrafast (<500
fs) energy transfer processes within the multimer, ‘slow’ (∼20 ps) energy transfer processes from peripheral RC
chlorophylls to the RC multimer, ultrafast charge separation (<500 fs) with a low yield starting from the singletexcited ‘accessory’ chlorophyll of the active branch, cation transfer from this ‘accessory’ chlorophyll to a ‘special
pair’ chlorophyll and/or charge separation starting from this ‘special pair’ chlorophyll (∼8 ps), and slow relaxation
(∼50 ps) of the radical pair by conformational changes of the protein. The charge separation in the PS II RC can
probably not be described as a simple trap-limited or diffusion-limited process, while for the PS II core and larger
complexes the transfer of the excitation energy to the PS II RC may be rate limiting.
Abbreviations: BRC – purple bacterial reaction center, Chl – chlorophyll, LHC II – light-harvesting complex
II, Phe – pheophytin, PS II – Photosystem II, P680 – primary electron donor of Photosystem II, QA – primary
plastoquinone electron acceptor, RC – reaction center, YZ – redox-active Tyr-161 of D1
Introduction
Photosystems are complex biological devices that are
able to convert visible light into an electrochemical
potential. Among all photosystems, Photosystem II
(PS II) is unique in the sense that it creates a much
stronger oxidant than other photosystems and that it
is even capable of oxidizing water to molecular oxygen. The green plant PS II is also one of the most
complex photosystems. It consists of more than 25 different protein subunits which work together to absorb
substantial amounts of visible light and transfer the
excitation energy to the photochemical reaction center
(RC). Within the RC, the electronically excited state of
a special chlorophyll (P680) induces the rapid transfer
of an electron to an adjacent pheophytin (Phe) molecule. The charge separation is then stabilized by the
electron transfer in about 200 ps from the reduced Phe
molecule to a plastoquinone molecule called QA , by
the electron transfer in nanoseconds from a redox active tyrosine (YZ ) at position 161 of the D1 protein to
P680+ (Diner and Babcock 1996), and by a number of
slower electron transfer reactions (millisecond timescale) which finally result in the oxidation of water to
molecular oxygen, the reduction of plastoquinone to
plastoquinol and the formation of a transmembrane pH
gradient. Other remarkable features of green plant PS
II include the very rapid turnover of one of the RC
proteins (the D1 protein) in plants under illumination
(Barber and Andersson 1992) and the direct involve-
196
Structure and organization
Figure 1. Model of the structural arrangement of the chlorin pigments in the PS II RC. The PS II RC probably contains ‘special pair’
and ‘accessory’ chlorophylls and pheophytins in similar positions
and orientations as in the RC of Rps. viridis or Rb. sphaeroides,
except that the ‘special pair’ chlorophylls are wider spaced and that
there are two ‘peripheral’ chlorophylls at a relatively large distance
from the six central chlorins.
ment of PS II in several regulatory mechanisms of the
photosynthesis (Bassi et al. 1997). A detailed understanding of the primary photosynthetic reactions in PS
II may, therefore, be important for the understanding
of photosynthetic energy conversion in general.
The primary reactions in PS II have recently been
reviewed by a number of groups (see, e.g. Van
Grondelle et al. 1994; Diner and Babcock 1996; Klug
et al. 1998). In this Minireview, we restrict ourselves
to a number of issues on the first reactions in PS
II that recently have been a subject of debate and
refer to the above-mentioned reviews for more detailed
overviews. The discussions include our views on the
suitability of the isolated PS II RC complex as a tool
to study primary charge separation in PS II, the origin of the 679 and 683 nm absorption bands of the
PS II RC complex, the origin of the relatively slow
charge separation kinetics in the PS II RC complex
at room temperature, the role and extent of excitonic
coupling in the reaction center and how this is realized
in the so-called multimer model (Durrant et al. 1995),
the equilibrium between the excited and radical pair
states and the mechanism of the primary charge separation reactions in PS II. The discussion is performed
in the light of the most recent structural data (Rhee
et al. 1998; Hankamer et al. 1999) and of new developments in the understanding of charge separation
in photosynthetic purple bacteria (Van Brederode and
Van Grondelle 1999).
PS II consists of at least 25 different types of protein
subunits, which are organized into two structurally and
functionally different parts (Hankamer et al. 1997).
The first part is the core complex, a well-defined
structure which is responsible for all electron transfer
reactions in PS II and which is organized as a dimer in the stacked, appressed regions of the thylakoid
membrane. The second part is the peripheral antenna,
which in green plants and algae consists of a collection
of light-harvesting complex II (LHC II) proteins and
which absorbs most of the light for PS II. A variable
part of the LHC II proteins is rather tightly associated
with the dimeric PS II core complex to form one of
the so-called PS II–LHC II super- or megacomplexes
(Boekema et al. 1999a). The structure of the trimeric
LHC II complex is known at 3.4 Å resolution, revealing the positions of 3 trans-membrane α-helices,
12 chlorophylls and 2 xanthophylls (Kühlbrandt et al.
1994), whereas the structure of the PS II core complex
without the CP43 core antenna protein is known at 8
Å resolution, revealing the positions of 23 transmembrane α-helices, of which 6 have been assigned to the
core antenna protein CP47, 10 to the reaction center
proteins D1 and D2 and 7 to small proteins consisting
of a single transmembrane α-helix (Rhee et al. 1998).
Near the central helices of the D1 and D2 proteins, a
number of masses were observed that were tentatively
assigned to 4 chlorophyll (Chl) and 2 pheophytin (Phe)
molecules, in line with the idea that the PS II RC binds
4 Chl a and 2 Phe a molecules in similar positions and
orientations as the corresponding molecules in the RC
of photosynthetic purple bacteria (see Figure 1 for a
schematic representation of the chlorophylls and pheophytins of the PS II RC). Also, the proposed wider
spacing between the chlorophylls of the ‘special pair’
(Braun et al. 1990; Durrant et al. 1995) became visible in the 8 Å map (Rhee et al. 1998). The D1 and
D2 proteins bind most probably two additional Chl a
molecules at conserved histidines at positions 118 and
117, respectively.
Most of the PS II protein complexes can now be
isolated and purified with at least partially retained
activity. For the PS II RC complex consisting of the
D1, D2, cytochrome b-559, PsbI and PsbW proteins,
several purification protocolls have been reported (see,
e.g. Eijckelhoff et al. 1996), but thus far, all preparations lack the quinone electron acceptors and the
possibility to perform secondary electron transfer. The
RC complex can also be isolated as a monomer with
197
two monomeric (CP29 and CP26) LHC II proteins
around the dimeric PS II core complex (Boekema et
al. 1999b). In the core complex, the positions are outlined of the D1 and D2 proteins, the CP47 and CP43
proteins and the small proteins of unknown function
(Hankamer et al. 1999). The most important features
are that S–LHC II can transfer excitation energy to
CP43 (either directly or via CP26 as an intermediate), that S–LHC II can also transfer excitation energy
to CP47 (via CP29 as a necessary intermediate), but
that there is no possibility of energy transfer between
the antenna proteins in the upper and lower parts of
the complex without reaching the D1 and D2 reaction
center proteins or the central part of the core dimer.
Spectroscopic properties
Absorption properties of the PS II RC
Figure 2. Projection map of the negatively stained ‘standard’ PS
II–LHC II supercomplex, obtained from the datasets reported in
Boekema et al. (1999a,b). In the PS II core parts, the regions in
red indicate the positions of the D1 and D2 proteins, while the ones
in green indicate the positions of the CP47 and CP43 core antenna
proteins and those in purple and yellow indicate the positions of
small proteins with one or two transmembrane α-helices (adopted
from Hankamer et al. 1999). The positions of trimeric (LHC II) and
monomeric (CP29 and CP26) peripheral antenna proteins are also
indicated in green to visualize the possible energy transfer pathways
to the D1 and D2 RC proteins. The structures on the two halves of
the PS II core dimer were slightly translationally shifted compared
to the data in Hankamer et al. (1999) to obtain an optimal overlay
with the PS II–LHC II supercomplex.
one (CP47) or two (CP47 and CP43) connected antenna proteins and partially retained activity and as
a dimer with even more connected antenna proteins
(the PS II–LHC II supercomplexes) and fully retained
activity (Schilstra et al. 1999).
Figure 2 shows a projection map of the most frequently observed PS II-LHC II supercomplex and
reveals the positions of one trimeric (S–LHC II) and
In contrast to the situation in the purple bacterial
reaction center (BRC), the Qy transitions of the chlorophylls and pheophytins of the PS II RC occur at about
the same wavelength (675 nm). This spectral congestion is one of the consequences of the weak coupling
between the various pigments (discussed in detail below) and considerably complicates the interpretation
of spectroscopic studies on the PS II RC. Only at
cryogenic temperatures some finestructure in the absorption spectrum can be observed with bands peaking
near 670 and 679 nm and a shoulder at about 683 nm
(Van Kan et al. 1990).
There is now good evidence that at least one of
the two peripheral chlorophylls absorbs maximally at
about 670 nm. In the 5 Chl PS II RC (RC-5) preparations, a chlorophyll peaking at 670 nm is missing
while the amplitude of the ‘slow’ (20–30 ps) energy
transfer from the peripheral to core chlorophylls is
about halved (Vacha et al. 1995). The second peripheral chlorophyll most likely also absorbs at 670
nm, because in the RC-5 preparations about half of
the slow 670 → 680 nm energy transfer process still
was observed (Vacha et al. 1995) and because the
remaining 4 K absorption around 670 nm in the RC5 preparations had exactly the same bandwidth and
peak wavelength as the total 4 K absorption around
670 nm in the RC-6 preparations (Eijckelhoff et al.
1997a). In the ‘red’ absorption region (679–683 nm),
it is well established that P680 and the ‘active’ pheophytin show significant contributions. There may also
be absorption from a ‘trap’ chlorophyll (Groot et al.
198
1996; Den Hartog et al. 1998a) responsible for the
trapping of excitations at very low temperatures (although this chlorophyll may also be part of P680 – see
below), and from the ‘inactive’ pheophytin molecule,
because chemical exchange of the inactive pheophytin
resulted in an absorbance decrease at about 680 nm
(Shkuropatov et al. 1997).
There is, however, no consensus yet on the absorption properties of the inactive pheophytin and very
recently Jankowiak et al. (1999) reported a very different peak wavelength at 668.3 nm. These authors
recorded a difference spectrum with a bleaching at
668.3 nm and a bandshift around ∼684 nm after addition of 4 mg/ml sodium dithionite in the dark in the
absence of oxygen and attributed this difference spectrum to reduction of the inactive pheophytin, because
the active pheophytin and the chlorophylls should
not be reduced under these conditions. Although the
theoretical basis of the assignment of the 668.3 nm
bleach to the inactive pheophytin seems reasonable,
the experimental evidence is weak (the bleaching of
the characteristic pheophytin Qx transition around 540
nm is obscured by the spectrum of the reduction of
cytochrome b-559). It is also not clear if possible pH
effects could have influenced the spectrum. Addition
of dithionite can easily lead to an acidification of the
medium, which can give rise to irreversible bleachings
at 684 and 665 nm (Yruela et al. 1999). In our view,
the interpretation of the results from the chemical exchange of the inactive pheophytin (Shkuropatov et al.
1997) is more straightforward and we, therefore, conclude that the inactive pheophytin most likely absorbs
around 680 nm.
There has also been quite some discussion and controversy in the literature on the origin of the 679 and
683 nm absorption bands. In some proposals, the 683
nm state was attributed to one of the two chlorophylls
located near the periphery of the complex and was
referred to as a ‘linker’ of excitation energy between
the core antenna chlorophylls and P680 (Seibert 1993;
Chang et al. 1994b). This interpretation is in conflict
with the above-mentioned idea that both peripheral
chlorophylls should absorb maximally near 670 nm.
In other reports, the 683 nm state was attributed to an
‘accessory’ chlorophyll in the central core of the PS II
RC (Konermann et al. 1997b), or to a special form of
P680 absorbing at long wavelength (Van der Vos et al.
1992; Kwa et al. 1994).
We note that the attribution of the 683 nm state
to an ‘accessory’ chlorophyll or to P680 does not
necessarily have to conflict, because the accessory
Figure 3. Line-narrowed emission spectra of PS II RC at 5 K.
With the technique of fluorescence line-narrowing (FLN), sub-nm
resolution emission spectra are recorded upon narrow-bandwidth
(∼cm−1 ) continuous-wave laser excitation (see, e.g. Peterman et
al. 1998). The aim of these FLN experiments is to record the
‘single-site’ emission spectra of the PS II RC absorbance bands
peaking at 680 and 684 nm. Generally, the single-site emission spectrum consists of a relatively narrow zero-phonon line (ZPL), caused
by the purely electronic transition, a broader wing, the so-called
phonon wing (PW), and a large number of less intense repeats of
the ZPL-PW structure at lower energy in emission. These repeats are
separated with the vibrational frequency from the purely electronic
transition, provide valuable information on the vibrational modes
of the emitting pigment(s) and can be used as a ‘fingerprint’ of the
emitting species, because the exact vibrational frequencies depend
on the molecular environment of the emitting species. Single-site
emission spectra can only be recorded if (a) the excitation is ‘selective’ (i.e. occurs by a laser), (b) the temperature is below ∼40
K and (c) energy transfer does not occur. The shown spectra are
the averages of six spectra selectively excited at 679.2 – 681.2 nm
(upper curves) and 682.8 – 686.0 nm (lower curves), each converted
to the wavenumber scale and plotted as the difference with the excitation wavenumber. The upper two spectra are 5 × magnifications
to enhance details in the vibronic region. Inset: magnification of the
region between 950 and 1350 cm−1 . The fact that line narrowing
is observed upon excitation at ∼680 nm confirms that the 679 and
683 nm spectral forms are not connected by energy transfer. The
vibrational finestructure in both spectra is very similar, which indicates that excitation of the 679 and 683 nm spectral forms leads to
fluorescence from chlorophyll molecules in the same protein environments. The differences in the low-frequency region arise from the
fact that the upper spectrum is excited mainly in the phonon side
band, whereas the lower spectrum is excited predominantly in the
zero-phonon lines.
chlorophyll may in fact be a part of P680 (see below).
Nevertheless, it is clear that the 679 and 683 nm absorption bands have both been observed in the states
directly responsible for charge separation at 4 K (radical pair and triplet – Van Kan et al. 1990), suggesting
that both absorption bands belong to P680. It has also
been shown that both spectral forms are not connected
by energy transfer at 4 K (Kwa et al. 1994), which
would be very unlikely if the 683 nm state would be
due to a peripheral chlorophyll.
199
However, not only the 4 K absorption data reveal two bands in the red part of the spectrum, also
the steady-state emission at 4 K shows two bands
peaking near 683.5 and 680.5 nm (Peterman et al.
1998). Figure 3 shows that selective excitation of the
chlorophylls absorbing around 680 and 684 nm gives
virtually identical vibrational finestructures of the 4 K
fluorescence, which strongly suggests that both emissions arise from the same molecule(s) or state(s) with
the same molecular environments. We stress that the
vibrational finestructures can be used as ‘fingerprints’
of the environment and that thusfar different finestructures were observed for different chlorophyll-protein
complexes. For the CP43 complex, it was even shown
that two different ‘red’ states in the same complex give
rise to different vibrational finestructures (Groot et al.
1999). These observations suggest that the two spectral forms have very similar origins and are the result
of an apparent ‘heterogeneity’.
We note that ‘heterogeneity’ has also been observed in the absorption band of the low-exciton component of the special pair chlorophylls of the reaction
center of photosynthetic purple bacteria (Hoff 1988;
Friesner and Won 1989) and that the heterogeneity
observed in the PS II RC does not have to be ‘real’.
It can be a direct result of the particular shape of the
‘single-site’ spectrum (the lower curve in Figure 3 –
see the legend of Figure 3 for an explanation of the
single-site spectrum). This spectrum shows on one
hand a very sharp zero-phonon line and a clear phonon
band at 19 cm−1 , and on the other a characteristic
and relatively intense feature around 80 cm−1 (see
also Peterman et al. 1998). The nonselective spectrum (the 4 K absorption spectrum) can be regarded
as a convolution of the single site spectrum with an
inhomogeneous distribution function, and if the inhomogeneous width is limited, it is expected that the
structures of (a) the zero-phonon line and 19 cm−1 feature and (b) the broad and relatively intense 80 cm−1
phonon side band will still be observed as separate
peaks. The fact that the energetic difference between
679 and 683 nm (85 cm−1 ) matches the energetic difference between these two structures is in line with this
idea.
Absorption properties of P680 in intact PS II
An important issue is the question to which extent the
spectroscopy of the purified PS II RC complex represents that of PS II in vivo. Based on experiments on
oxygen-evolving PS II membranes from higher plants
and PS II preparations from cyanobacteria, it has been
suggested that in intact PS II the primary electron
donor has only one absorption band at 683–684 nm
at cryogenic temperatures, that increased biochemical
manipulation of the photosystem leads to a shift to
679–680 nm, and thus that the 679 nm state could be
non-physiological (Hillmann et al. 1995). While this
suggestion is in line with the idea that the 679 nm
and 683 nm spectral forms arise from the same (set
of) pigment(s), it is remarkable that PS II RC particles
prepared by very different methods (long, short or no
Triton X-100 incubation, the 5 Chl preparation) all
contain about the same ratio of the 683 and 679 nm
spectral forms and that increased biochemical manipulation of purified reaction centers only seems to result
in a broadening of the absorption bands (Eijckelhoff et
al. 1996, 1997a).
We would like to point out that if intact PS II contains two (or more) spectral forms of P680, the ones
peaking near 680 nm could remain undetected at 4 K.
In other words, it is possible that only the red states
could give rise to charge separation at 4 K. The argument is that in intact PS II many excitations will reach
the long-wavelength states of the CP47 and CP43 core
antenna proteins absorbing at 683–690 nm (Chang et
al. 1994a; Groot et al. 1999), which at 4 K are generally too low in energy to allow uphill energy transfer
to the 680 nm state of P680. Thus, in complexes with a
blue RC state a large part of the excitation energy will
be trapped on the low-energy states of CP47 and CP43,
while in complexes with a red RC state, many excitations will be transferred to the RC and, therefore, give
rise to charge separation. In both cases (blue or red RC
state), a ∼684 nm state will be bleached upon cation
or triplet formation, in line with the experimental results (Hillmann et al. 1995). This possibility gives a
straightforward explanation for the observed blueshift
of the bleaching to 680 nm upon raising the temperature in intact PS II (Hillmann et al. 1995), because
at higher temperatures the uphill energy transfer to the
blue RC states will become possible. We conclude that
there is no evidence that the absorption properties of
the chlorins in isolated PS II RC particles differ from
those in intact PS II. Conversely, it should be kept in
mind that the quinone electron acceptors are missing
in isolated PS II RC complexes, which has a strong
influence on the electron transfer reactions within this
complex.
200
Cation, triplet and fluorescing states of the PS II RC
At low temperatures, the triplet state of the PS II RC is
localized on a single Chl with its plane tilted 30◦ relative to the membrane (Van Mieghem et al. 1991). This
orientation is almost identical to that of the accessory
chlorophylls in the BRC (Kwa et al. 1994). The tripletminus-singlet difference spectrum is characterized by
a bleaching of the 131 C=O stretch mode at 1669 cm−1
(Noguchi et al. 1993).
At higher temperatures, the situation becomes
more complicated. Between 100 and 200 K, the spectrum of the triplet changes from that of the abovementioned single species with its plane tilted 30◦
relative to the membrane to that of two different species in which the species observed at low-temperature
is now mixed with a second species with its plane vertical to the membrane plane (Kamlowski et al. 1996).
The latter orientation is similar to that of the ‘special
pair’ in the BRC.
The emission at 4 K (Peterman et al. 1998) gives
rise to a very similar vibrational finestructure as observed for the triplet, such as the 131 C=O stretch mode
at 1669 cm−1 and a characteristic low-frequency mode
at 80 cm−1 (Kwa et al. 1994). These and other observations suggest that the steady-state emission and the
triplet arise from the same species. Between 70 and
150 K, the shape of the emission spectrum changes
from a spectrum peaking at 684 nm to a spectrum
peaking at 682 nm (Groot et al. 1994), suggesting
that also here a different species starts to contribute
at higher temperatures.
The vibrational finestructure of the P680+ -minusP680 difference spectrum has been recorded at 150 K
(Noguchi et al. 1998) and shows features of charge delocalization. The 131 C=O stretch mode is under these
circumstances characterized by bleachings at 1679 and
1704 cm−1 . These energies differ from those observed
for the triplet difference spectrum at 85 K, although it
is not clear to which extent the much higher temperature gives rise to these differences. Nevertheless, the
different finestructures have been used as an argument
in favour of triplet migration to a Chl different from the
species responsible for charge separation (Noguchi et
al. 1998). In our view, there is no clear evidence for
such a migration. Apart from the temperature problem mentioned above, it can not be excluded that the
observed frequencies of vibrational modes vary as a
function of experimental conditions. The presence of
chemicals to produce the radical pair state could influence the charge distribution around the secondary
donors, which could influence the frequencies of the
131 C=O stretch modes. The absence of triplet migration would in every case explain the highly polarized
triplet-minus-singlet absorbance-difference spectrum
obtained upon 4 K excitation at λ > 680 nm (Kwa et
al. 1994).
Excitonic coupling in PS II
When two or more pigments are sufficiently close
in space, they will interact and, consequently, the
transition energies, orientations and relative intensities of the original transition dipoles will be modified
(see, e.g. Van Grondelle et al. 1994; Van Amerongen et al. 2000). This excitonic interaction is optimal
when the frequencies of the involved transitions are
the same and weakens progressively upon increasing
the difference between these frequencies.
Experimental evidence for a relatively weak excitonic interaction (∼140 cm−1 ) in the PS II RC has
been presented by Kwa et al. (1994), who detected
differently-oriented low- and high-exciton components at 680–683 and 667 nm, respectively, and by
Chang et al. (1994b), who reported a high-exciton
component at 667 nm on the basis of hole-burning
experiments. This coupling most probably occurs
between the chlorins of the primary donor P680, and
is much weaker than the coupling between the primary
donor pigments in the BRC of Rb. sphaeroides
(∼550 cm−1 ) or Rps. viridis (∼950 cm−1 ). There has
been a debate on the extent of excitonic interactions
between the other chlorins in the central core part of
the PS II RC. Some authors argued that these interactions can be neglected and that P680 should be viewed
as a weakly-coupled dimer of chlorophylls (see, e.g.
Bosch et al. 1995; Konermann and Holzwarth 1996),
whereas others argued that the interactions between all
six chlorins should be taken into account (Tetenkin et
al. 1989; Durrant et al. 1995) and that the central part
of the PS II RC should be viewed as a weakly-coupled
multimer of four Chl a and two Phe a molecules.
The interaction energy or coupling strength V12
between two interacting identical pigments can be
calculated using the point-dipole approximation:
V 12 = cκ
D
|R312 |
in which V12 is given in cm−1 , c is a constant (5.04
in theory and about 5.3 in practice – Kwa 1993),
κ is a dimensionless geometrical factor that can be
201
Table 1. Relative angles between calculated exciton bands of a trimer consisting of
two Chl a and one Phe a molecule coordinated as the PM , BL , and HL molecules of
the Rps. viridis RC, respectively (see Figure 1). Also listed are the dipole strengths
in Debye2 . The simple point-dipole approximation was used (see text), and the
monomer wavelengths were taken to be 674 nm for all three pigments. The dipole
strengths of Chl a and Phe a were assumed to be 23 and 15 Debye2 , while c was
assumed to be 5.3
Exciton
band
λ (nm)
Dipole
strength
(Debye2 )
Angle with
band 1
(degrees)
Angle with
band 2
(degrees)
Angle with
band 3
(degrees)
1
2
3
680.5
673.2
668.4
49.3 (82%)
8.0 (13%)
3.1 (5%)
–
68.3
80.9
68.3
–
49.2
80.9
49.2
–
calculated from the crystal structure, D is the dipole
strength in (Debye)2 and R̄12 is the vector connecting
the dipoles (in nm). The point-dipole approximation
has been used successfully to map the excitonic interactions in structurally well-resolved FMO complexes
from green sulphur bacteria (Louwe et al. 1997).
Kwa (1993) used the point-dipole approximation
to calculate the extent of excitonic coupling in a trimer consisting of one Phe a and two Chl a molecules
positioned exactly as the HL , BL and PM molecules
of the Rps. viridis RC (see also Table 1) and found
three exciton bands with similar energies, intensities
and orientations to those observed experimentally in
PS II RC complexes. Calculations using a complete
RC structure have suggested that the interaction energy between the two ‘special pair’ chlorophylls is a
very critical parameter (Durrant et al. 1995). In a situation as in the bacterial reaction center, this interaction
is so strong that a special pair dimer is formed with
very minor contributions from the other RC pigments
and with one major red-shifted low-exciton band. The
other extreme is a situation in which this interaction
is negligible, such as in the situation as in Table 1. In
this situation, there are two groups of three chlorins
which give rise to two sets of exciton bands, each delocalized over separate ‘arms’ of the reaction center,
and thus to two nearly degenerate low-exciton bands.
It is not clear which situation occurs in the PS II RC.
Some observations suggest the presence of two nearly
degenerate exciton bands (see below), suggesting a
situation of a very small interaction energy between
the two ‘special pair’ chlorophylls, but an intermediary situation of equal interaction energy between all
chlorins (and thus of a ‘multimer’ of maximally six
interacting chlorins) is also very well possible.
The calculations by Kwa (1993) refer to a situation of degenerate site energies. This is, however,
not very realistic, for two different reasons. First, the
average site energies of pigments at different sites
in a pigment–protein complex may not be the same
(Gudowska-Nowak et al. 1990). Second, there is a
spread of site energies of pigments at the same site
in the complete ensemble of molecules (the inhomogeneous width or energetic disorder). An energetic
disorder of the order of about 100 cm−1 was taken into
account in the calculations presented by Durrant et al.
(1995). This number may be higher if exchange narrowing of the absorption band has occurred. However,
there are, as yet, no indications for large differences
in the average site energies of the constituting chlorins. Reduction of the active pheophytin, chemical
exchange of the inactive pheophytin molecule, or the
formation of a triplet, either at very low temperatures
on a chlorophyll oriented as the ‘accessory’ bacteriochlorophyll in the BRC, or at higher temperatures on a
chlorophyll oriented as a ‘special pair’ bacteriochlorophyll in the BRC, results in all cases in an absorbance
decrease around 680 nm (see above), which would not
be expected if one of these components would have a
very different average site energy.
We conclude from these considerations that excitonic interactions between the six central chlorins
of the PS II RC should not be neglected. An additional argument for this can be found in the most
recent structural data (Rhee et al. 1998), which seem
to suggest that the mutual distances between the six
central chlorins, except those of the special pair, do
not differ very much in the PS II RC and BRC and
that the distance parameters in the exciton calculations
mentioned above must have been estimated about cor-
202
rectly. Thus, a situation arises in the PS II RC in which
the coupling between the chlorins has about the same
value as the intrinsic disorder. Consequently, a multimer model arises in which there is no ‘special pair’
and in which for each realization of the disorder, the
excitation may be dynamically localized on basically
any combination of neighbouring chlorins. It is not
difficult to imagine that at sufficiently low temperatures such a system shows a very complex behaviour,
and that large variations of transition energies and dipole strengths can be expected from RC to RC (see,
e.g. Figures 2 and 3 in Durrant et al. 1995).
We note that the multimeric organization of the
central chlorins of the PS II RC (and of any other
organization of nearby chlorophylls) implies that the
redox potentials of all constituting chlorophylls must
be very high, as discussed recently by Mulkidjanian
(1999). In this view, both the electrostatic influence
of Arg-181 of the D2 subunit and a retarded protonic
relaxation near the central chlorins contribute to the
raise of the redox potential of the oxidized chlorophyll
to up to about 1.15 V.
Kinetics of energy transfer and trapping in the PS
II RC
The kinetics of energy transfer and trapping in the PS
II RC have been studied by several groups using both
ultrafast absorbance-difference and fluorescence techniques. After intense debates in the literature, it seems
now generally accepted that primary charge separation
at room temperature is a strongly multiphasic process with main components of about 8, 20 and 50 ps
(see, e.g. Müller et al. 1996; Greenfield et al. 1997;
Donovan et al. 1997; Klug et al. 1998). Thus, the
formation of the primary radical pair takes more time
in the isolated PS II RC than in the BRC, where most
components are shorter than 10 ps. Excitation energy
transfer within the six central core pigments has been
reported to occur in maximally 500 fs (Durrant et al.
1992; Merry et al. 1996), whereas the energy transfer
from the peripheral chlorophylls to the central core occurs in about 20–30 ps (Roelofs et al. 1993; Schelvis
et al. 1994; Rech et al. 1994).
Several origins have been proposed for the relatively slow charge separation kinetics in the PS II
RC: 1) Delocalization of the excitation energy (Van
Grondelle et al. 1994). In case of trap-limited kinetics
(i.e. when the equilibration of the excitation energy is
much faster than charge separation), the rate of charge
separation is proportional to the probability to find the
excitation on the primary electron donor. This probability is 1/N, where N is the number of pigments
among which the excitation energy can be divided. In
the PS II RC N equals 6–8, depending on the extent
to which the two peripheral chlorophylls contribute to
the equilibration of the excitation energy. In the BRC
N equals 1, because only the lowest excitonic state of
the special pair contributes to the 870 nm absorption
band and the 800 and 770 nm states of the accessory chlorophylls and the pheophytins are too high in
energy to become populated. 2) Slow energy transfer
(Groot et al. 1997). It has been suggested on the basis
of low-temperature experiments that the PS II RC contains pigments or states degenerate with P680 (the
so-called ‘trap’-states, which trap excitation energy at
cryogenic temperatures – Groot et al. 1996), that the
energy transfer between the P680 and trap states is
rate-limiting at intermediate temperatures, and that energy transfer may also be limiting at room temperature
(Groot et al. 1997). This latter suggestion, however,
seems at variance with the conclusion of Merry et al.
(1996) and Leegwater et al. (1997) that the energy
transfer between such states is about 500 fs at room
temperature, which is too fast to be rate-limiting. 3)
An initially about zero free energy difference of charge
separation combined with slow charge-induced relaxations in the protein surroundings (Konermann et al.
1997a). These authors concluded that charge separation to the initial radical pair state is actually very fast
in the PS II RC, that the yield of this first radical pair
state is rather low because the equilibrium between the
excited states and the radical pair is towards the excited states, and that subsequent energetic relaxations
in the protein surroundings, induced by the creation
of the two charges, shift this equilibrium towards the
radical pair, ultimately resulting in a high yield of radical pair states and, apparently, relatively slow charge
separation kinetics. An about zero free-energy difference of the primary charge separation reaction of PS
II was also concluded from quantum chemical calculations using a density-functional theory (Blomberg et
al. 1998).
A consequence of the latter explanation is that the
intrinsic rate of charge separation must be very fast,
perhaps even in the order of a few hundred femtoseconds, and perhaps even faster than the rate of
excitation equilibration in the central core part of the
PS II RC. In the BRC, similar ultrafast rates have
already been recorded for charge separation from the
excited ‘accessory’ bacteriochlorophyll on the active
203
branch (Van Brederode et al. 1997, 1999). In intact
bacteria, the importance of this very fast charge separation route is limited because most excitation energy
will end up in the bacteriochlorophylls of the LH1
antenna, which are too low in energy to excite the
accessory bacteriochlorophyll. In PS II, however, the
energetic differences between the various chlorophylls
are negligible at room temperature, and a significant
part of the excitation energy can become localized
on the ‘accessory’ chlorophylls. The delocalization of
the excitation energy (the first possibility mentioned
above) probably plays a significant role as well. It
should, however, be realized that within the PS II RC
the description in terms of trap-limited kinetics may
not be appropriate in view of the possible faster rate of
charge separation than excited state equilibration.
If in the first charge separated state the electron
hole is on the ‘accessory’ chlorophyll of the active
branch, then electron transfer from one of the ‘special pair’ chlorophylls to this hole may occur, and if
the free energy of the latter charge separated state is
lower than the former, as is usually the case in bacterial
reaction centers (Van Brederode and Van Grondelle
1999), then this electron transfer will shift the equilibrium between the excited state and the radical pair
towards the radical pair, thus providing a stabilization
of the charge separation reaction. A good candidate
for this process is the ∼8 ps phase, which has been
observed in both transient absorption (Müller et al.
1996; Greenfield et al. 1997) and ultrafast fluorescence (F. van Mourik et al., unpublished observations).
Slow energy transfer reactions (mainly involving the
‘peripheral’ chlorophylls) and protein relaxations may
then be primarily responsible for the ∼20 and 50 ps
processes, respectively.
In contrast to the situation at room temperature,
only a few studies have been performed on the kinetics of charge separation at cryogenic temperatures.
However, even from the limited available evidence it
is clear that also the temperature dependence of charge
separation differs considerably in RCs of PS II and
purple bacteria. In the latter RC, charge separation occurs in about 3 ps at room temperature and in about
1 ps at 8 K (Fleming et al. 1988). In the PS II RC,
however, the kinetics have been shown to slow down
upon cooling. Data between 240 K and 20 K could
best be described by phases of 0.4 ps and 18 ps at 240
K that progressively retard to 2.6 ps and 120 ps at 20
K (Groot et al. 1997), while at 7 K phases of about
5 and 120 ps were observed (Greenfield et al. 1999).
The main implication of these results is that charge
separation is activationless in the BRC and at least to
some extent activated in the PS II RC.
The situation of an initially about zero free energy
difference of charge separation can have large effects
on the efficiency of charge separation at very low temperatures. In such a situation, the charge separation
reaction will have a small negative 1G in some RC
complexes and a small positive 1G in others (depending on the energetic disorder of the system). In other
words, charge separation will proceed nearly activationless and fast in some complexes, but activated and
(very) slow in others, because the thermal energy will
not be sufficient to activate the reaction at very low
temperatures. The latter situation may lead to a longlived primary electron donor unable to perform charge
separation at very low temperatures, thus explaining
a number of very similar properties of P680 and the
species responsible for the fluorescence at very low
temperatures (Peterman et al. 1998). It is, therefore,
very well possible that the chlorins responsible for
the ‘trap’ of excitation energy at very low temperatures (Groot et al. 1994, 1996) and for P680 are the
same and that the difference is only determined by the
precise value of the free energy difference of charge
separation.
The excited-state radical pair equilibrium in PS II
It is now beyond doubt that a dynamic equilibrium
between the primary radical pair and chlorophyll excited states plays a very important role in the charge
separation reaction in PS II RC complexes. A nice
example was reported by Merry et al. (1998), who
analyzed a set of isolated PS II RC complexes from
site-directed mutants of the cyanobacterium Synechocystis PCC 6803, and found that a modulation of the
free energy of the radical pair by changes in the direct
environment of the P680+ or Phe− states correlates
linearly with the equilibrium constant of the charge
separation reaction and the quantum yield of charge
separation. The free energy change of the charge separation reaction was estimated to be –27 meV for PS
II RC complexes from green plants at a time of 60
ps after the initiation of the charge separation reaction
(Merry et al. 1998). This number is probably less negative at earlier times and more negative at later times
due to dynamic relaxations of the radical pair. We
note that there may also be considerable variation in
the free energy values at fixed times due to structural
heterogeneities (Groot et al. 1994).
204
The influence of the equilibrium between the
chlorophyll excited states and the radical pair in larger
systems (such as PS II core complexes or membranes)
is more difficult to establish. The exciton/radical
pair equilibrium model, originally proposed by Van
Grondelle (1985) and Van Gorkom (1985), has been
used successfully to explain the time-resolved fluorescence kinetics in PS II core particles (Schatz et al.
1988), in particular because it provides a good explanation for the biphasic decay in ‘open’ centers (with
oxidized quinone acceptor QA ). In this view, the first
phase with about 50–100 ps kinetics reflects the trapping of the excitation energy in the primary radical
pair P680+ Phe− , while the second phase with about
300–500 ps kinetics reflects the electron transfer from
to Phe− toQA ,which indeed has been shown to proceed
with these kinetics (see, e.g. Nuijs et al. 1986).
There are a number of indications that the situation is much more complex than originally proposed
by Schatz et al. (1988): (1) a 250–300 ps phase has
also been detected in PS II core particles with doubly
reduced QA (Van Mieghem et al. 1992), which indicates that this phase can not be attributed solely to
charge stabilization; (2) a non-radiative decay channel
has been proposed in the PS II RC with kinetics of,
again, about 300 ps (Merry et al. 1998), and it can
not be excluded that this process is also important for
the 300–500 ps kinetics in PS II core particles; (3)
the assumption of Schatz et al. (1988) of a fast (<
10 ps) equilibration in the PS II core antenna may
not be correct. The most important hint for this is
the 8 Å structure of the CP47–RC complex (Rhee et
al. 1998), which seems to point to a relatively large
distance between the groups of chlorophylls in the
CP47 and RC parts of the complex, although it cannot be excluded yet that both groups of chlorophylls
are connected by one or more linker chlorophylls not
observed in the 8 Å structure. The helices I and II
of the D1 and D2 proteins, however, seem to not be
located between CP47 and the central part of the RC,
which suggests that if the ‘peripheral’ chlorophylls of
the PS II RC are really located near these helices, as
is expected from their slow (∼ 20 ps) energy transfer to the central chlorins, then they probably do not
function as ‘linkers’ of excitation energy transfer from
core antenna to RC. Moreover, there is now conclusive evidence that the CP47 and CP43 core antenna
proteins are located at opposite sites from the PS II
RC (see, e.g. Eijckelhoff et al. 1997b; Harrer et al.
1998; Rhee et al. 1998) and that, therefore, direct excitation energy transfer between these antenna systems
is impossible. All these new structural details point
to a relatively large distance between core antenna
and RC, a situation which is also encountered in PS
I and purple bacterial photosystems (Van Grondelle et
al. 1994), which favours a situation of transfer-to-thetrap-limited kinetics (Beekman et al. 1994; Valkunas
et al. 1995), and which suggests that the 50–100 ps
trapping phase may to at least some extent be determined by a slow transfer of excitation energy from the
CP43 or CP47 core antenna to the PS II RC.
We would furthermore like to mention that experiments on PS II membranes or complete thylakoids
are very hard to interpret in view of the very heterogeneous organization of the peripheral antenna LHC
II in these systems (see, e.g. Boekema et al. 1999
and Dekker et al. 1999). The observed trapping times
of 80, 200 and 390 ps in ‘open’ PS II membranes
(Vass et al. 1993) may relate to PS II centers with
increasing amounts of LHC II, while minor phases
with lifetimes of 2–3 ns may reflect the fluorescence
properties of LHC II trimers or multimers that are not
directly connected to the PS II core. Fluorescence lifetime measurements on carefully prepared PS II–LHC
II supercomplexes and LHC II multimers will shed
more light on this issue.
Conclusions
In Figures 4 and 5 we show schematic models that may
account for the most elementary energy transfer and
charge separation mechanisms at physiological temperatures discussed above for the PS II RC and the
PS II core, respectively. The most essential features
of the model of the processes in the PS II RC (Figure 4) are: (A) energy transfer on two levels: ultrafast
population transfer of delocalized exciton states within
the six central chlorins of the RC (with time constants
of ∼100 and ∼500 fs), and slow (∼20 ps) energy
transfer from the peripheral chlorophylls to the RC
core, (B) ultrafast charge separation with a low yield,
primarily starting from the singlet-excited ‘accessory’
Chl on the active branch, (C) cation transfer from the
‘accessory’ Chl to a ‘special pair’ Chl and/or charge
separation starting from the ‘special pair’ Chl on the
active branch (∼8 ps), and (D) slow relaxation (∼50
ps) of the radical pair by conformational changes of
the protein. At very low temperatures, the triplet delocalization does not occur, indicating that process (C)
does not occur to a significant extent, but now part of
the energy transfer and/or initial charge separation re-
205
Figure 5. Schematic representation of the energy transfer and
primary charge separation reactions in a monomeric PS II core
complex. For details see text.
Figure 4. Schematic representation of the energy transfer and
primary charge separation reactions in the PS II RC. For details see
text.
actions may slow down very considerably. According
to our model, both processes proceed between almost isoenergetic states, which in view of the disorder
means that they can either be slightly downhill (and
thus proceed fast) or slightly uphill (and thus become
retarded very significantly at very low temperatures).
The most essential point of the model in Figure 5 is
a possibly slow energy transfer from the core antenna
to the PS II RC. We note that all lifetimes depicted in
both models are very rough estimates of the real values, and that at room temperature some energy transfer
routes may proceed in the same time-range as charge
separation. The charge separation in the PS II RC can,
therefore, probably not be described as a simple traplimited or diffusion-limited process, while for the PS II
core and larger complexes the transfer of the excitation
energy to the PS II RC may be rate limiting.
206
Acknowledgements
We thank Drs Egbert Boekema and Erwin Peterman
for providing data and help with Figures 2 and 3,
respectively. Our research was supported by the Netherlands Foundation for Scientific Research (NWO) via
the Foundation for Physical Research (FOM) and the
Foundation for Life and Earth Sciences (ALW).
References
Barber J and Andersson B (1992) Too much of a good thing: Light
can be bad for photosynthesis. Trends Biochem Sci 17: 61–66
Bassi R, Sandonà D and Croce R (1997) Novel aspects of chlorophyll a/b proteins. Physiol Plant 100: 769–779
Beekman LMP, Van Mourik F, Jones MR, Visser HM, Hunter CN
and Van Grondelle R (1994) Trapping kinetics in mutants of the
photosynthetic purple bacterium Rhodobacter sphaeroides: Influence of the charge separation rate and consequences for the
rate-limiting step in the light-harvesting process. Biochemistry
33: 3143–3147
Blomberg MRA, Siegbahn PER and Babcock GT (1998) Modeling
electron transfer in biochemistry: A quantum chemical study of
charge separation in Rhodobacter sphaeroides and Photosystem
II. J Am Chem Soc 120: 8812–8824
Boekema EJ, Van Roon H, Calkoen F, Bassi R and Dekker JP
(1999a) Multiple types of association of Photosystem II and its
light-harvesting antenna in partially solubilized Photosystem II
membranes. Biochemistry 38: 2233–2239
Boekema EJ, Van Roon H, Van Breemen JFL and Dekker JP
(1999b) Supramolecular organization of Photosystem II and its
light-harvesting antenna in partially solubilized Photosystem II
membranes. Eur J Biochem 266: 444–452
Bosch MK, Proskuryakov II, Gast P and Hoff AJ (1995) Relative
orientation of the optical transition dipole and triplet axes of the
Photosystem II primary donor. A magnetophoto-selection study.
J Phys Chem 99: 15310–15316
Braun P, Greenberg BM and Scherz A (1990) D1-D2-cytochrome
b-559 complex from the aquatic plant Spirodela oligorrhiza:
Correlation between complex integrity, spectroscopic properties,
photochemical activity and pigment composition. Biochemistry
29: 10376–10387
Chang H-C, Jankowiak R, Yocum CF, Picorel R, Alfonso M, Seibert
M and Small GJ (1994a) Exciton level structure and dynamics in
the CP47 antenna complex of Photosystem II. J Phys Chem 98:
7717–7724
Chang H-C, Jankowiak R, Reddy NRS, Yocum CF, Picorel R,
Seibert M and Small GJ (1994b) On the question of the chlorophyll a content of the Photosystem II reaction center. J Phys
Chem 98: 7725–7735
Dekker JP, Van Roon H and Boekema EJ (1999) Heptameric association of light-harvesting complex II trimers in partially solubilized Photosystem II membranes. FEBS Letters 449: 211–214
Den Hartog FTH, Vacha F, Lock AJ, Barber J, Dekker JP and Völker
S (1998a) Comparison of the excited-state dynamics of five- and
six-chlorophyll Photosystem II reaction center complexes. J Phys
Chem B 102: 9174–9180
Diner BA and Babcock GT (1996) Structure, dynamics, and energy conversion efficiency in Photosystem II. In: Ort DR and
Yocum CF (eds) Oxygenic Photosynthesis. The Light Reac-
tions, pp 213–247. Kluwer Academic Publishers, Dordrecht, The
Netherlands
Donovan B, Walker LA II, Kaplan D, Bouvier M, Yocum CF and
Sension RJ (1997) Structure and function in the isolated reaction center complex of Photosystem II. 1. Ultrafast fluorescence
measurements of PS II. J Phys Chem B 101: 5232–5238
Durrant JR, Hastings G, Joseph DM, Barber J, Porter G and Klug
DR (1992) Subpicosecond equilibration of excitation energy in
isolated Photosystem II reaction centers. Proc Natl Acad Sci
USA 89: 11632–11636
Durrant JR, Klug DR, Kwa SLS, Van Grondelle R, Porter G and
Dekker JP (1995) A multimer model for P680, the primary
electron donor of Photosystem II. Proc Natl Acad Sci USA 92:
4798–4802
Eijckelhoff C, Van Roon H, Groot M-L, Van Grondelle R and
Dekker JP (1996) Purification and spectroscopic characterization of Photosystem II reaction center complexes isolated with
or without Triton X-100. Biochemistry 35: 12864–12872
Eijckelhoff C, Vacha F, Van Grondelle R, Dekker JP and Barber J
(1997a) Spectroscopic characterization of a 5 Chl a Photosystem
II reaction center complex. Biochim Biophys Acta 1318: 266–
274
Eijckelhoff C, Dekker JP and Boekema EJ (1997b) Characterization
by electron microscopy of dimeric Photosystem II core complexes from spinach with and without CP43. Biochim Biophys
Acta 1321: 10–20
Fleming GR, Martin JL and Breton J (1988) Rates of primary
electron transfer in photosynthetic reaction centers and their
mechanistic implications. Nature 333: 190–192
Friesner RA and Won Y (1989) Spectroscopy and electron transfer dynamics of the bacterial photosynthetic reaction center.
Biochim Biophys Acta 977: 99–122
Greenfield SR, Seibert M, Govindjee and Wasielewski MR (1997)
Direct measurement of the effective rate constant for primary
charge separation in isolated Photosystem II reaction centers. J
Phys Chem B 101: 2251–2255
Greenfield SR, Seibert M and Wasielewski MR (1999) Timeresolved absorption changes of the pheophytin Qx band in isolated Photosystem II reaction centers at 7 K: Energy transfer and
charge separation. J Phys Chem B 103: 8364–8374
Groot M-L, Peterman EJG, Van Kan PJM, Van Stokkum IHM,
Dekker JP and Van Grondelle R (1994) Temperature dependent
triplet and fluorescence quantum yields of the Photosystem II
reaction center described in a thermodynamic model. Biophys J
67: 318–330
Groot M-L, Dekker JP, Van Grondelle R, Den Hartog FTH and
Völker S (1996) Energy transfer and trapping in isolated Photosystem II reaction centers of green plants at low temperature. A
study by spectral hole-burning. J Phys Chem 100: 11488–11495
Groot M-L, Van Mourik F, Eijckelhoff C, Van Stokkum IHM,
Dekker JP and Van Grondelle R (1997) Charge separation in
the reaction center of Photosystem II studied as function of
temperature. Proc Natl Acad Sci USA 94: 4389–4394
Groot M-L, Frese RN, de Weerd FL, Bromek K, Pettersson Å, Peterman EJG, Van Stokkum IHM, Van Grondelle R and Dekker JP
(1999) Spectroscopic properties of the CP43 core antenna protein
of Photosystem II. Biophys J 77: 3328–3340
Gudowska-Nowak E, Newton MD and Fajer J (1990) Conformational and environmental effects on bacteriochlorophyll optical
spectra: Correlations of calculated spectra with structural results.
J Phys Chem 94: 5795–5801
Hankamer B, Barber J and Boekema EJ (1997) Structure and membrane organization of Photosystem II from green plants. Ann Rev
Plant Phys Plant Mol Biol 48: 641–672
207
Hankamer B, Morris EP and Barber J (1999) Revealing the structure of the oxygen-evolving core dimer of Photosystem II by
cryoelectron crystallography. Nature Struct Biol 6: 560–564
Harrer R, Bassi R, Testi MG and Schäfer C (1998) Nearestneighbour analysis of a Photosystem II complex from
Marchantia polymorpha L. (liverwort), which contains reaction
centre and antenna proteins. Eur J Biochem 255: 196–205
Hillmann B, Brettel K, Van Mieghem F, Kamlowski A, Rutherford
AW and Schlodder E (1995) Charge recombination reactions in
Photosystem II. 2. Transient absorbance difference spectra and
their temperature dependence. Biochemistry 34: 4814–4827
Hoff AJ (1988) Bacterial reaction centers are intrinsically heterogeneous. In: Breton J and Verméglio A (eds) The Photosynthetic
Bacterial Reaction Center, Structure and Dynamics, pp 89–97.
Plenum Press, New York
Jankowiak R, Rãtsep M, Picorel R, Seibert M and Small GJ (1999)
Excited states of the 5-chlorophyll Photosystem II reaction
center. J Phys Chem B 103: 9759–9769
Jennings RC, Bassi R, Garlaschi FM, Dainese P and Zucchelli
G (1993) Distribution of the chlorophyll spectral forms in the
chlorophyll–protein complexes of Photosystem II antenna. Biochemistry 32: 3203–3210
Kamlowski A, Frankemüller L, Van der Est A, Stehlik D and
Holzwarth AR (1996) Evidence for delocalization of the triplet
state 3 P680 in the D1 D2 cytb559-complex of Photosystem II. Ber
Bunsenges Phys Chem 100: 2045–2051
Klug DR, Durrant JR and Barber J (1998) The entanglement of
excitation energy transfer and electron transfer in the reaction
centre of Photosystem II. Phil Trans R Soc 356: 449–464
Konermann L and Holzwarth AR (1996) Analysis of the absorption
spectrum of Photosystem II reaction centers: Temperature dependence, pigment assignment and inhomogeneous broadening.
Biochemistry 35: 829–842
Konermann L, Gatzen G and Holzwarth AR (1997a) Primary processes and structure of the Photosystem II reaction center. 5.
Modeling of the fluorescence kinetics of the D1–D2-cyt-b559
complex at 77 K. J Phys Chem B 101: 2933–2944
Konermann L, Yruela I and Holzwarth AR (1997b) Pigment assignment in the absorption spectrum of the Photosystem II reaction
center by site-selected fluorescence spectroscopy. Biochemistry
36: 7498–7502
Kühlbrandt W, Wang DN and Fujiyoshi Y (1994) Atomic model
of plant light-harvesting complex by electron crystallography.
Nature 367: 614–621
Kwa SLS (1993) Plant solar cells under illumination. A spectroscopic investigation of Photosystem II. Doctoral Thesis, Vrije
Universiteit Amsterdam
Kwa SLS, Eijckelhoff C, Van Grondelle R and Dekker JP (1994)
Site-selection spectroscopy of the reaction center complex of
Photosystem II. I. Triplet-minus-singlet absorption difference:
A search for a second exciton band of P-680. J Phys Chem 98:
7702–7711
Leegwater JA, Durrant JR and Klug DR (1997) Exciton equilibration induced by phonons: Theory and application to PS II
reaction centers. J Phys Chem B 101: 7205–7210
Louwe RJW, Vrieze J, Hoff AJ and Aartsma TJ (1997) Towards
an integral interpretation of the optical steady-state spectra of
the FMO-complex of Prosthecochloris aestuarii. 2. Exciton
simulations. J Phys Chem B 101: 11280–11287
Merry SAP, Kumazaki S, Tachibana Y, Joseph DM, Porter G,
Yoshihara K, Barber J, Durrant JR and Klug DR (1996) Subpicosecond equilibration of excitation energy in isolated Photosystem II reaction center revisited: Time-dependent anisotropy.
J Phys Chem 100: 10469–10478
Merry SAP, Nixon PJ, Barter LMC, Schilstra M, Porter G, Barber J,
Durrant JR and Klug DR (1998) Modulation of quantum yield of
primary radical pair formation in Photosystem II by site-directed
mutagenesis affecting radical cations and anions. Biochemistry
37: 17439–17447
Mulkidjanian AY (1999) Photosystem II of green plants: On the
possible role of retarded protonic relaxation in water oxidation.
Biochim Biophys Acta 1410: 1–6
Müller MG, Hucke M, Reus M and Holzwarth AR (1996) Primary
processes and structure of the Photosystem II reaction center. 4.
Low-intensity femtosecond transient absorption spectra of D1D2-cyt-b559 reaction centers. J Phys Chem 100: 9527–9536
Noguchi T, Inoue Y and Satoh K (1993) FTIR studies on the
triplet state of P680 in the Photosystem II reaction center:
Triplet equilibrium within a chlorophyll dimer. Biochemistry 32:
7186–7195
Noguchi T, Tomo T and Inoue Y (1998) Fourier transform infrared
study of the cation radical of P680 in the Photosystem II reaction
center: Evidence for charge delocalization on the chlorophyll
dimer. Biochemistry 37: 13614–13625
Nuijs AM, Van Gorkom HJ, Plijter JJ and Duysens LNM (1986)
Primary charge separation and excitation of chlorophyll a in
Photosystem II particles from spinach as studied by picosecond
absorbance difference spectroscopy. Biochim Biophys Acta 848:
167–175
Peterman EJG, Van Amerongen H, Van Grondelle R and Dekker JP
(1998) The nature of the excited state of the reaction center of
Photosystem II of green plants. A high-resolution fluorescence
spectroscopy study. Proc Natl Acad Sci USA 95: 6128–6133
Rech T, Durrant JR, Joseph MD, Barber J, Porter G and Klug DR
(1994) Does slow energy transfer limit the observed time constant for radial pair formation in photosystem II reaction centers?
Biochemistry 33: 14768–14774
Rhee K-H, Morris EP, Barber J and Kühlbrandt W (1998) Threedimensional structure of the plant Photosystem II reaction centre
at 8 Å resolution. Nature 396: 283–286
Roelofs TA, Kwa SLS, Van Grondelle R, Dekker JP and Holzwarth
AR (1993) Primary processes and structure of the Photosystem
II reaction center. II. Low temperature picosecond fluorescence
kinetics of a D1–D2-Cytochrome b-559 reaction center complex
isolated by short Triton-exposure. Biochim Biophys Acta 1143:
147–157
Satoh K (1996) Introduction to the Photosystem II reaction center – isolation and biochemical and biophysical characterization.
In Ort DR and Yocum CF (eds) Oxygenic Photosynthesis: The
Light Reactions, pp 193–211. Kluwer Academic Publishers,
Dordrecht, The Netherlands
Schatz GH, Brock H and Holzwarth AR (1988) A kinetic and energetic model for the primary processes in Photosystem II. Biophys
J 54: 397–405
Schelvis JPM, Van Noort PI, Aartsma TJ and Van Gorkom HJ
(1994) Energy transfer, charge separation and pigment arrangement in the reaction center of Photosystem II. Biochim Biophys
Acta 1184: 242–250
Schilstra MJ, Nield J, Dörner W, Hankamer B, Carradus M, Barter
LMC, Barber J and Klug DR (1999) Similarity between electron donor side reactions in the solubilized Photosystem II-LHC
II supercomplex and Photosystem II-containing membranes.
Photosynth Res 60: 191–198
Seibert M (1993) Biochemical, biophysical, and structural characterization of the isolated Photosystem II reaction center complex. In: Deisenhofer J and Norris JR (eds) The Photosynthetic
Reaction Center, Vol I, pp 319–356. Academic Press, New York
208
Shkuropatov AY, Khatypov RA, Volshchukova TS, Shkuropatova
VA, Owens TG and Shuvalov VA (1997) Spectral and photochemical properties of borohydride-treated D1–D2-cytochrome
b-559 complex of Photosystem II. FEBS Lett 420: 171–174
Tetenkin VL, Gulyaev BA, Seibert M and Rubin AB (1989) Spectral
properties of stabilized D1/D2/cytochrome b-559 Photosystem II
reaction center complex. FEBS Lett 250: 459–463
Vacha F, Joseph DM, Durrant JR, Telfer A, Klug DR, Porter G
and Barber J (1995) Photochemistry and spectroscopy of a fivechlorophyll reaction center of Photosystem II isolated by using a
Cu affinity column. Proc Natl Acad Sci USA 92: 2929–2933
Valkunas L, Liuolia V, Dekker JP and Van Grondelle R (1995) Description of energy migration and trapping in Photosystem I. The
application of a model with two scaling parameters. Photosynth
Res 43: 149–154
Van Amerongen H, Valkunas L and Van Grondelle R (2000)
Photosynthetic Excitons. World Scientific, Singapore
Van Brederode ME and Van Grondelle R (1999) New and unexpected pathways for ultrafast electron transfer in photosynthetic
reaction centers. FEBS Lett 455: 1–7
Van Brederode ME, Jones MR, Van Mourik F, Van Stokkum IHM
and Van Grondelle R (1997) A new pathway for transmembrane electron transfer in photosynthetic reaction centers of
Rhodobacter sphaeroides not involving the excited special pair.
Biochemistry 36: 6855–6861
Van Brederode ME, Van Mourik F, Van Stokkum IHM, Jones MR
and Van Grondelle R (1999) Multiple pathways for ultrafast
transduction of light energy in the photosynthetic reaction center of Rhodobacter sphaeroides. Proc Natl Acad Sci USA 96:
2054–2059
Van der Vos R, Van Leeuwen PJ, Braun P and Hoff AJ (1992) Analysis of the optical absorbance spectra of D1–D2-cytochrome
b-559 complexes by absorbance-detected magnetic resonance.
Structural properties of P680. Biochim. Biophys. Acta 1140:
184–198
Van Gorkom (1985) Electron transfer in Photosystem II. Photosynth
Res 6: 97–112
Van Grondelle (1985) Excitation energy transfer, trapping and annihilation in photosynthetic systems. Biochim Biophys Acta 811:
147–195
Van Grondelle R, Dekker JP, Gillbro T and Sundström V (1994) Energy transfer and trapping in photosynthesis. Biochim Biophys
Acta 1187: 1–65
Van Kan PJM, Otte SCM, Kleinherenbrink FAM, Nieveen MC,
Aartsma TJ and Van Gorkom HJ (1990) Time-resolved spectroscopy at 10 K of the Photosystem II reaction center; deconvolution of the red absorption band. Biochim Biophys Acta 1020:
146–152
Van Mieghem FJE, Satoh K and Rutherford AW (1991) A chlorophyll tilted 30◦ relative to the membrane in the Photosystem II
reaction centre. Biochim Biophys Acta 1058: 379–385
Van Mieghem FJE, Searle GFW, Rutherford AW and Schaafsma
TJ (1992) The influence of the double reduction of QA on the
fluorescence decay kinetics of Photosystem II. Biochim Biophys
Acta 1100: 198–206
Vass I, Gatzen G and Holzwarth AR (1993) Picosecond timeresolved fluorescence studies on photoinhibition and double
reduction of QA in Photosystem II. Biochim Biophys Acta 1183:
388–396
Yruela I, Tomas R, Alfonso M and Picorel R (1999) Effect of the pH
on the absorption spectrum of the isolated D1–D2-cytochrome
b559 complex of Photosystem II. J Photochem Photobiol B 50:
129–136