Vol.:(0123456789)
1 3
Photochemical & Photobiological Sciences
https://doi.org/10.1007/s43630-023-00362-z
ORIGINAL PAPERS
Ultrafast protein response inthePfr state ofCph1 phytochrome
YangYang1· TillStensitzki1· ChristinaLang3· JonHughes3· MariaAndreaMroginski2· KarstenHeyne1
Received: 17 September 2022 / Accepted: 27 December 2022
© The Author(s) 2023
Abstract
Photoisomerization is a fundamental process in several classes of photoreceptors. Phytochromes sense red and far-red light
in their Pr and Pfr states, respectively. Upon light absorption, these states react via individual photoreactions to the other
state. Cph1 phytochrome shows a photoisomerization of its phycocyanobilin (PCB) chromophore in the Pfr state with a time
constant of 0.7ps. The dynamics of the PCB chromophore has been described, but whether or not the apoprotein exhibits an
ultrafast response too, is not known. Here, we compare the photoreaction of 13C/15N labeled apoprotein with unlabeled apo-
protein to unravel ultrafast apoprotein dynamics in Cph1. In the spectral range from 1750 to 1620 cm−1 we assigned several
signals due to ultrafast apoprotein dynamics. A bleaching signal at 1724 cm−1 is tentatively assigned to deprotonation of a
carboxylic acid, probably Asp207, and signals around 1670 cm−1 are assigned to amide I vibrations of the capping helix close
to the chromophore. These signals remain after photoisomerization. The apoprotein dynamics appear upon photoexcitation
or concomitant with chromophore isomerization. Thus, apoprotein dynamics occur prior to and after photoisomerization on
an ultrafast time-scale. We discuss the origin of the ultrafast apoprotein response with the ‘Coulomb hammer’ mechanism,
i.e. an impulsive change of electric field and Coulombic force around the chromophore upon excitation.
Graphical abstract
1 Introduction
Phytochromes are photoreceptors of plants, bacteria and
fungi that use red and far-red light as a source of informa-
tion regarding their environment. Light absorption by a bilin
chromophore initiates a reaction cascade that finally regu-
lates fundamental processes, such as photosynthesis, flower-
ing, seed germination, and shade avoidance. Phytochromes
act as a bistable photoswitch between the Pr (red absorbing)
state and the Pfr (far-red absorbing) state. Photoexcitation
of the lowest energy Pr state triggers photoisomerization
* Karsten Heyne
karsten.he[email protected]
1 Department ofPhysics, Freie Universität Berlin, Arnimallee
14, 14195Berlin, Germany
2 Institut Für Chemie, Technische Universität Berlin, Straße
des 17. Juni 135, 10623Berlin, Germany
3 Institut für Pflanzenphysiologie, Justus-Liebig Universität
Giessen, Senckenbergstr. 3, 35390Giessen, Germany
Photochemical & Photobiological Sciences
1 3
of the bilin chromophore and formation of the first ground-
state intermediate Lumi-R on a picosecond time scale, fol-
lowed by relaxation via Meta-R intermediates to form the
Pfr state. Photoexcitation of Pfr, on the other hand, leads to
sub-picosecond isomerization and Lumi-F production fol-
lowed by Meta-F intermediates prior to reformation of Pr
[1–15]. How these photoisomerizations are brought about
is still unclear, however.
The crystal structure of the photosensory module of the
cyanobacterial phytochrome Cph1 as Pr [16] like that of
plant phytochromes [17], shows a remarkable overall dumb-
bell structure. The N-terminal PAS-GAF bidomain is tied
together by a figure-8 knot, the GAF domain enveloping
the chromophore while the PHY domain is held at a dis-
tance via a long α-helix. However, a remarkable hairpin loop
known as the tongue extends back from the PHY domain
to make contact with the GAF domain, thereby sealing the
chromophore pocket. It has been suggested that refolding of
the tongue pulls on the PHY domain, movement of which
activates the physiological signal [18]. Since a crystal struc-
ture of Cph1 as Pfr is not yet available, a structural model
(Fig.1a) was built by homology modeling based on the Pfr
crystal structure of a distantly-related bacteriophytochrome
[19, 20]. This model was used to identify ultrafast chromo-
phore changes during Pfr photoisomerization and heteroge-
neity in the Pfr ground state [20]. In this study, we compare
our published data of 13C/15N labeled apoprotein [20, 21]
with the measured data of unlabeled apoprotein, and use the
Pfr model (see Fig.1) as a starting point to identify ultrafast
protein changes upon photoexcitation.
In general, it is assumed that photoisomerization of the
chromophore is the decisive reaction step initiating all sub-
sequent structural changes in the photoreceptor from pico-
seconds to seconds. This assumption is supported by several
investigations using chemically modified chromophores that
are unable to photoisomerize (‘locked’ chromophores) and
consequently fail to form the photoproduct state, e.g. Pfr
[22]. Furthermore, ultrafast investigations on photorecep-
tors with ‘locked’ chromophores exhibit dynamics on the
picosecond to nanosecond time scale, but their impact on
the formation of an active state remains elusive [1, 23–27].
Photoexcitation can result in largely altered dipole
moments in the electronic excited state. A significant dipole
moment increase was observed in the electronically excited
state of retinal bound proteins, and the created electric fields
were proposed to interact with the protein [28, 29]. The
Fig. 1 a Homology model (βf or down form) of the Pfr state of Cph1
phytochrome [20] with the phycocyanobilin (PCB) chromophore and
its electronic transition dipole moment (tdm) represented by a green
arrow. b enlarged view into the binding pocket of the chromophore
PCB; the blue colored α-helical part was called “capping helix” in
DrBphP [37]; here the backbones of arg254, ser206, and backbone
of side chains 259–266 are colored in blue, these backbone carbonyl
groups exhibit vtdms with a relative angle between 14° and 60° with
the electronic transition dipole moment (green vector). Other back-
bone carbonyl groups are oriented mainly perpendicular to the elec-
tronic tdm or are far away from PCB. c enlarged view on the PCB
chromophore in the down conformation [20] and arrangement of
important amino acids, such as Asp207 (D207)
Photochemical & Photobiological Sciences
1 3
altered electric field at the chromophore was proposed to
be the driving force for its isomerization and interact with
charged and polarizable apoprotein groups in its near envi-
ronment [28, 29]. The energy absorbed by the retinal could
be used for new bond arrangements in the apoprotein struc-
ture affecting proton positions and stabilizing retinal con-
formations [30]. The interaction with the altered dipole was
expected to be on the time-scale of nanoseconds or longer
to induce relevant conformational apoprotein changes [29].
These hypotheses could not be confirmed by direct experi-
mental data for a long time. Gross etal. demonstrated apo-
protein changes of the amide I band of bR in the electronic
excited state of a locked retinal persisting up to the nanosec-
ond time scale [23]. Later on energy relaxation to the apo-
protein was demonstrated in other proteins after isomeriza-
tion [31, 32]. However, the impact of these observations on
downstream processes of the photocycle remained elusive.
Recently, ultrafast structural changes on apoprotein side
chains were reported prior to and after isomerization of
the chromophore in bR, CaChR 1, and Agp2 phytochrome
[33–35] and have been connected to downstream processes
leading to the photoactive state [34, 35]. The underlying
mechanism was suggested to be a ‘Coulomb hammer’-
like process rapidly changing the electric field around the
chromophore upon its photoexcitation, thereby triggering
changes in hydrogen bond networks and protonation states
[36]. Ultrafast X-ray crystallography measurements were
reported for a bacteriophytochrome from Deinococcus
radiodurans (DrBphP) in its Pr state at 1ps and 10ps after
excitation [37]. The structural alterations at 1ps show a twist
of the D-ring, the dissociation of the pyrrole water from
the chromophore, movement of ring A and a key aspartate
(D207), as well as ultrafast water and backbone movements
around the chromophore. Significant backbone movements
in DrBphP were located at the “capping helix” (blue colored
structural motif in Fig.1b) [37]. Ultrafast water movement
around the chromophore as well as apoprotein changes
were reported also for bR by ultrafast X-ray crystallogra-
phy [38, 39]. Compared to other ultrafast methods, X-ray
crystallography reports a huge number of structural rear-
rangements throughout the apoprotein. However, since these
experiments were performed with very high pump intensi-
ties, it is unclear whether the structural changes occur in
a single photon excitation regime or are a consequence of
multi-photon processes [39]. Clarifying whether substantial
structural changes prior to and after isomerization occur in
photoreceptors on the ultrafast time-scale is fundamental to
understanding apoprotein dynamics.
Comparison of X-ray results for the Pr state of DrBphP
with other methods at single photon excitation intensities
with Cph1 reveals contradictory results for the twist of the
D-ring, and movement of ring A [40, 41]. Whether these
differences are due to the different phytochromes or methods
used is unclear. As contributions of the isomerizing chromo-
phore dominate the vibrational signals, very little is known
about ultrafast changes in the apoprotein upon photoexcita-
tion. To address this, we compared ultrafast, polarization-
resolved VIS pump—IR probe experiments in the Pfr state
of Cph1 with and without 13C/15N labelled apoprotein. By
comparing the dynamics in each case, we have been able to
identify ultrafast apoprotein changes in the spectral region
between 1610 cm−1 and 1750 cm−1 not only after but also
prior to photoisomerization.
2 Materials andmethods
Preparation of the 13C/15N labeled apoprotein Cph1Δ2 (non-
labelled PCB chromophore) was described previously [20,
21, 42–44]. Unlabelled holophytochrome was generated via
invitro autoassembly of apoCph1Δ2 with PCB extracted
by methanolysis of phycobiliproteins from commercially
available Spirulina pellets dissolved in 100mM potassium
phosphate buffer (pH 7.0) following procedures described
earlier [45]. The SEC-purified sample was rebuffered into
D2O-TESß at pD 7.8 by three passes through a home-built
pressure cell. The samples had an optical density of 0.15 OD
to 0.4 OD at 710nm.
We use polarization resolved femtosecond visible (VIS)
pump infrared (IR) probe spectroscopy to directly investi-
gate the dynamics of the photoreaction. Measurements on
13C/15N labeled apoprotein were described elsewhere [20].
Measurements on unlabeled holophytochrome were per-
formed in a similar way: pump and probe pulses were gen-
erated using home built optical parametric amplifiers. By
difference frequency mixing in various steps, we obtained
tunable mid-IR probe pulses of 300fs (FWHM) or shorter
at a repetition rate of 1.088kHz. Simultaneously, laser
pulses of ~ 150fs duration around 720nm with a bandwidth
(FWHM) of ~ 10nm were generated and used to initiate the
Pfr photoreaction (system response ~ 350fs). At this wave-
length no Pr absorption occurs. A 640nm laser diode was
used for the background illumination to enrich the Pfr state.
Photoselection experiments were performed by overlap-
ping one pump beam with a diameter of 300μm, and two
probe beams with diameters of 150μm on the sample. The
pulse energy of the pump and probe was 0.55μJ and 10 nJ,
respectively. The excitation efficiency was 15% or less. After
passing the sample, both probe pulses were dispersed with
an imaging spectrograph and recorded with a 2 × 32 element
MCT array detector (resolution ∼1.5 cm−1). The polarization
direction of the pump pulse was changed after every scan
by a half-wave (λ/2) plate, to switch the relative polariza-
tions between pump and two probe beams from parallel to
perpendicular or perpendicular to parallel. Thus, we detect
absorption signal for parallel Apar and perpendicular Aper
Photochemical & Photobiological Sciences
1 3
polarization. The signals for isotropic polarization Aiso were
taken from Aiso = (Apar + 2 × Aper)/3. The global analysis was
performed on the complete dataset with three shared time
constants and an offset. Error margins represent 1σ devia-
tions (68.27%). An exhaustive search analysis with a broader
error margin of 3σ (99.7%) provides time constants with
asymmetric error margins of τ1 = 0.3–0.7ps, τ2 = 2.1–5.5ps,
and τ3 = 9.6–18ps.
Samples were placed between two 1mm thick CaF2 win-
dows of 38mm diameter in a sample holder with a 50µm
Teflon spacer. The sample was moved continuously in hori-
zontal and vertical direction by a Lissajous-scanner to ensure
sample exchange between two successive pump pulses. All
experiments were performed at room temperature. The opti-
cal density at 640nm was between 0.3 and 0.6 when the
sample was fully in the Pr state. The Pr-Pfr-Pr photocycle
was tested before each experiment to ensure that the samples
were photoactive.
Polarization resolved measurements use photoselection
to induce a transient orientation of excited molecules in the
sample. With respect to the pump pulse polarization, the
probe pulse is detected for parallel and perpendicular polari-
zations under identical conditions. The dichroic ratio D is
defined as the ratio of parallel versus perpendicular polar-
ized absorption signal. The dichroic ratio is a measure for
the relative angle between the excited electronic transition
dipole moment (tdm) and the detected vibrational transi-
tion dipole moment (vtdm). Since the electronic tdm is fixed
within the excited molecule, the relative angle can provide
orientational information on the detected vibrational group
within the protein [40]. The electronic tdm of the S0 → S1
transition of the Pfr state is shown as a green arrow in Fig.1.
2.1 Vibrational frequencies andtransition dipole
moments
We used the published “down conformation” model from
Stensitzki etal. [20] as a structural model of Cph1 in the
Pfr state. In this structure, the Asp207 is modeled with an
anionic COO group. In addition to it, a structural model was
generated with a protonated Asp207 by manually protonat-
ing the carboxylic side chain. The DFT/CHARMM opti-
mized structures of PCB were used as input for frequency
calculations and computation of electronic and vibrational
transition dipole moments. In all these calculations the pro-
tein environment is described as a cloud of atomic point
charges from the CHARMM force field. The procedure is
described in detail [46, 47]. Tdm’s of electronic transitions
were computed by time-dependent (TD)-DFT theory imple-
mented in Gaussian09 software [48]. The relative angle
between the electronic tdm and specific vtdm’s can be esti-
mated directly from these calculations.
3 Results
Femtosecond polarization resolved VIS-pump IR-probe
experiments were performed upon excitation of the Pfr state
of Cph1 in D2O around 720nm and probing in the spec-
tral range from 1750 to 1610 cm−1. In this spectral range
carbonyl vibrations are dominant and are easily identified
by 13C labeling. For example, the high frequency C = O
stretching vibration of a carboxyl group at 1730 cm−1 shifts
upon 13C labeling by about 42 cm−1 to ~ 1690 cm−1. Amide
I vibrations of a backbone C = O group at 1669 cm−1 shift
upon 13C labeling to frequencies of 1626 cm−1. The 15N
labeling has only minor influence on the spectral position
of the amide I group. Other vibrational groups in this spec-
tral range are high frequency CN vibrations of arginine
[49]. Upon 13C/15N labeling a red-shift of about 30 cm−1
is observed and a red-shift of about 70 cm−1 is reported in
D2O [50–52].
3.1 Absorbance difference spectra
In Fig.2a and 2c absorbance difference spectra of Cph1
phytochrome are presented at different pump—probe delay
times upon photoexcitation for unlabeled and 13C/15N
labeled apoprotein, respectively. The PCB chromophore is
unlabeled and we expect identical chromophore dynamics in
both measurements. Indeed, the dynamics are very similar
(see Figs.2a and 2c), although distinct alterations are visible
around 1724 cm−1, 1680 cm−1, and 1635 cm−1. Differences
in the spectra can be attributed to the 13C/15N labeling of the
apoprotein, resulting in a frequency down-shift of apoprotein
vibrations. Polarization resolved averaged absorbance differ-
ence spectra for parallel, perpendicular and isotropic probe
polarization with respect to pump pulse polarization are
shown in Fig.2b and 2d for unlabeled and 13C/15N labeled
apoprotein, respectively. Upon photoexcitation, the vibra-
tional dynamics of the 13C/15N labeled apoprotein show time
constants of 0.7 ± 0.3ps and 5 ± 2ps in the spectral range
from 1750 to 1610 cm−1 [20], where apoprotein contribu-
tions are expected to be negligible due to 13C/15N labeling.
We averaged in the time window from 10 to 50ps, after the
ultrafast chromophore dynamics are mainly completed [21].
3.2 Dynamics of13C/15N labeled protein
Absorbance difference spectra at selected pump-probe
delay times are presented in Fig.2c for isotropic probe
polarization. Dynamics of the Pfr state of Cph1 with
13C/15N labeled apoprotein were investigated and discussed
in detail elsewhere [20, 21]. In short, the difference spectra
show a strong negative (bleaching) signal at 1708 cm−1
Photochemical & Photobiological Sciences
1 3
with a small shoulder at 1702 cm−1, as well as a negative
signal at ~ 1724 cm−1 at 0.5ps delay time. These signals at
1724 cm−1 and around 1705 cm−1 appear with the system
response, decay with 0.7 ± 0.2ps and 5 ± 2ps, and were
assigned to C = O carbonyl stretching vibrations of PCB
rings A and ring D, respectively. The characteristic band
of the Pfr photoisomerization is the vibrational band of the
ring D carbonyl stretching vibration. Its negative bleach-
ing band is located at 1708 cm−1 and decays to about 20%
of its initial value at 50ps after photoexcitation [21]. The
remaining bleaching signal fraction is a measure of the
quantum yield of the photoisomerization. The small shoul-
der at 1702 cm−1 was also assigned to the C = O stretching
vibration of ring D representing a different ground-state
conformation and demonstrating heterogeneity of the elec-
tronic ground state of PCB [9, 20, 53]. At longer delay
times, the bleaching band at 1724 cm−1 decays to zero and
is superimposed upon the rise of a positive signal assigned
to the carbonyl C = O stretching absorption of ring D in
the Lumi-F photoproduct [21]. The photoisomerization
of the chromophore is completed at 30ps with a posi-
tive signal at 1724 cm−1 reflecting the isomerized PCB in
Lumi-F, and negative signal at 1708 cm−1 from the ini-
tial Pfr ground state bleaching. These two bands show an
absorption strength peak ratio of about 1:1 as expected
for the same C = O stretching vibration before and after
photoisomerization [21]. After 30ps no other signals are
detected in the spectral range from 1750 to 1620 cm−1 in
13C/15N labeled Cph1 apoprotein (Fig.2c).
Polarization resolved absorbance difference spectra
averaged from 10 to 50ps are presented in Fig.2d around
1720 cm−1. The positive signal at 1724 cm−1 reflecting
Lumi-F absorption show very similar absorption strengths
for parallel (black dots) and perpendicular (open red circles)
polarizations. The dichroic ratio D = Apar/Aper reflects the rel-
ative angle between the excited electronic tdm (see Fig.1)
and the probed vtdm at a given wavenumber [24, 54]. The
relative angle θ can range between 0° and 90° and can be
determined from the dichroic ratio D by θ = arccos([(2D–1)/
(D + 2)]½). The dichroic ratio can be directly extracted
from the parallel and perpendicular polarized absorbance
difference spectra at a given delay time and wavenumber.
From Fig.2d the dichroic ratio for the Lumi-F absorption
DLF at 1724 cm−1 and the ring D bleaching signal DBL at
1708 cm−1 can be directly estimated to DLF = 1.3 (θ = 46°)
and DBL = 1.5 (θ = 41°), respectively. These values agree
with dichroic ratios of DLF = 1.3 ± 0.2 (θ between 41° and
51°) and DBL = 2.0 ± 0.5 (θ between 19° and 41°) determined
Fig. 2 Absorption difference spectra of Cph1 in D2O upon photo-
excitation around 720nm at different pump-probe delay times. The
PCB chromophore is unlabeled. Left: isotropic polarized absorption
spectra for unlabeled protein (a) and 13C/15N labeled apoprotein (c).
Right: scaled polarization resolved absorption spectra averaged from
10 to 50ps delay time in the spectral window around 1720 cm−1. Par-
allel (black spheres), perpendicular (red open circles), and isotropic
(purple line) polarizations with respect to pump pulse polarization for
unlabeled apoprotein (c), and 13C/15N labeled apoprotein (d). Aver-
aged data in (c) are scaled by a factor of two. Differences between
labeled and unlabeled apoprotein are clearly visible. Additional pro-
tein contributions shift the apparent band positions at early delay
times; after photoreaction at 50 ps the position of the peaks (grey
lines) are identical at the given resolution. The dataset on the 13C/15N
labeled apoprotein was discussed elsewhere [20]
Photochemical & Photobiological Sciences
1 3
by simulating the polarization resolved absorption spectra
[20].
3.3 Unlabeled protein dynamics
The absorbance difference spectra of the unlabeled apo-
protein are dominated by the negative bleaching signal at
1708 cm−1 as presented in Fig.2a. This bleaching band
reflects mainly the dynamics of the carbonyl group of ring
D in the parent state. Comparison of the bleaching signal
strength upon photoexcitation and after photoexcitation
reflects the portion of the excited molecules relaxing back
to its parent state and the portion undergoing forward reac-
tion. Similar to the photoreaction of the 13C/15N labeled apo-
protein, the bleaching signal decreases 30% between delay
times of 0.5ps to 50ps (see Fig.2a and 2c). Moreover,
the bleaching transients depicted in Fig.3b indicate nearly
identical behavior. Thus, we can conclude that the ultrafast
isomerization dynamics of PCB are identical in labeled and
unlabeled apoprotein with a forward quantum yield of about
20%.
The missing contribution of the ring D carbonyl bleach-
ing signal at 1708 cm−1 is converted to a new positive
signal of ring D carbonyl stretching vibration in the Lumi-
F conformation. This positive product signal exhibits a
characteristic absorption at 1724 cm−1. At this frequency
position, a bleaching signal due to carbonyl stretching
absorption of ring A dominates the dynamics at 0.5ps
in Fig.2a and 2c. This bleaching contribution decays to
zero on a few picoseconds. Within a picosecond Lumi-
F is formed and its positive contribution dominates for
longer delay times. At around 50ps the positive signature
of Lumi-F at 1724 cm−1 is expected to have similar abso-
lute strength compared to the missing contribution of the
ring D carbonyl bleaching signal at 1708 cm−1 as reported
for labeled apoprotein and shown in Fig.2c. In contrast, in
the unlabeled apoprotein the positive signal at 1724 cm−1
is nearly absent at 50ps delay time (Fig.2a). Thus, we
expect an additional negative apoprotein contribution
around 1724 cm−1 masking the positive contribution of
the PCB at longer delay times. Comparison of the tran-
sients at 1724 cm−1 for labeled and unlabeled apoprotein
are presented in Fig.3a. The difference between unlabeled
(green line) and labeled (blue line) transients gives a ris-
ing negative signal within a picosecond that stays constant
up to 50ps. This negative signal reflects the contribu-
tion of unlabeled apoprotein. The frequency position at
1724 cm−1 is characteristic of C = O stretching vibrations
of carboxylic acid and carbonyl groups. From Fig.2b
the dichroic ratio for the ring D bleaching signal, DBL,
at 1708 cm−1 can be directly estimated as ~ 1.3 (θ = 46°),
matching the value derived for labeled apoprotein within
the error margins. In contrast, the DLF at 1724 cm−1 devi-
ates strongly from the findings in labeled apoprotein. The
Fig. 3 Isotropic polarized tran-
sients of unlabeled (blue dots),
and 13C/15N labelled apoprotein
(green circles) are presented on
a linear and logarithmic time
scale together with simulated
transients (blue and green
lines). Differences induced by
apoprotein labeling are directly
visible by comparing the blue
and green lines. Isotropic polar-
ized transients at the frequency
position of the Lumi-F marker
band, and the system response
(grey dotted line) (a) and the
ground-state marker band (b)
are displayed. Polarization
resolved transients for parallel
(black lines) and perpendicular
(red lines) polarization together
with isotropic polarized tran-
sients (green and blue dots) are
presented at 1682 cm−1 (c) and
1630 cm−1 (d)
Photochemical & Photobiological Sciences
1 3
perpendicular polarized signal is much smaller compared
to the parallel one (Fig.2b). This can be explained by a
negative signal with a stronger contribution for perpen-
dicular than parallel polarization. This means the dichroic
ratio D < 1 corresponding to a relative angle of the vtdm to
the excited electronic tdm between 54° and 90°.
A broad positive signal around 1680 cm−1 is detected
upon photoexcitation (Fig.2a). The positive contribution at
1690 cm−1 with a smaller shoulder from 1680 to 1660 cm−1
was attributed to the ring D carbonyl stretching signal in
the electronic excited state in labeled apoprotein [20]. In
the unlabeled apoprotein the positive contribution around
1680 cm−1 is broader and more pronounced. We assign this
positive feature to the dynamics of chromophore’s ring D
and apoprotein dynamics. In addition, a negative signal
of similar strength is observed at 1650 cm−1, both signals
decaying on a similar time-scale of about 10ps. This signa-
ture could reflect amide I dynamics. Around 1630 cm−1 a
broad negative signal decays on a time-scale of about 10ps.
The transient is shown in Fig.3d.
Differences between the labeled (red lines) and unlabeled
(black lines) protein dynamics can also be identified by com-
paring their absorbance difference spectra in Fig.4a. The
spectra are scaled to the same bleaching signal at 1708 cm−1
at a delay time of 1ps. At this delay time notable differences
are found around 1680 cm−1 and 1640 cm−1 with stronger
positive and negative signals in the unlabeled apoprotein,
respectively. These differences become smaller at delay
times of 15ps and 50ps, and only subtle deviations are vis-
ible in this spectral range at 50ps. In contrast, differences at
1724 cm−1 show up at 15ps and increase up to 50ps. Fur-
thermore, small differences around 1700 cm−1 are apparent
at 15ps and 50ps. This indicates apoprotein contributions
in addition to chromophore dynamics around 1724 cm−1 and
1708 cm−1.
A global analysis of the dynamics of the unlabeled
protein provides four components with time constants of
0.4 ± 0.1ps, 3.5 ± 0.6ps, and 12 ± 2ps, and a constant com-
ponent. The decay associated spectra (DAS) are presented
in Fig.4b. The shortest time component of 0.4ps (not
shown) is affected by our system response of about 0.35ps
and reflects a mixture of the isomerization time constant of
0.7ps and nonlinear signals due to temporal pump-probe
overlap. The DAS for 3.5ps in unlabeled apoprotein shows
similar spectra features compared to the reported DAS for
5.0ps in labeled apoprotein [20]. Thus, the 3.5ps compo-
nent mainly reflects chromophore relaxation dynamics. This
is expected, since the chromophore contributes strongest to
the IR difference signal due to its direct excitation.
In the investigated spectral range, we found no dynamics
of the 13C/15N labeled apoprotein, since the contributions
are down-shifted out of the observed spectral window. In the
unlabeled apoprotein, we found an additional time constant
of 12ps that we assign to apoprotein dynamics. We expect
to find the same time constant at lower frequencies in the
labeled apoprotein.
Upon photoexcitation, the PCB chromophore is pro-
moted to its electronic excited state. A part of the popula-
tion isomerizes with 0.7ps and forms Lumi-F, while the
remaining part decays back to Pfr with time constants of
0.7ps and ~ 5ps [20]. Excited state signatures of the PCB
vibrations are clearly visible at 1695 cm−1 with a declin-
ing flank down to 1650 cm−1 (in Fig.2b). These features
are gone at 15ps delay time. In parallel to the PCB signals
apoprotein contributions appear within our time resolution
persisting longer than 50ps. Observed apoprotein dynamics
Fig. 4 Absorbance difference spectra of Pfr upon excitation around
720nm at selected delay times of 1ps, 15ps, and 50ps. a for 13C/15N
labeled apoprotein (red lines) and unlabeled apoprotein (black lines);
b Decay associated spectra of the dynamics of unlabeled protein for
isotropic polarization; for the unlabeled protein an additional time
constant of 12 ps was found; the “const” time constant indicates
that dynamics do not change on the time-scale of hundreds of pico-
seconds; it reflects contributions from Lumi-F (positive) and ground
state (negative); the spectra for 0.4ps is not shown due to interfer-
ences with the system response
Photochemical & Photobiological Sciences
1 3
show decaying negative signals at 1724 cm−1, 1708 cm−1,
1660 cm−1, and 1640 to 1600 cm−1, while around 1690 cm−1
we found a positive decaying signal. We tentatively assign
the positive and negative signals around 1690 cm−1 and
1660 cm−1, respectively, to an amide I signal of the pro-
tein backbone close to the chromophore. The backbone car-
bonyl groups of the “capping helix” (see Fig.1b, 1c, and 1d)
exhibit a carbonyl stretching vtdm between 14° and 60°. The
average impact on several carbonyl groups of the “capping
helix” should show a dichroic ratio of about 1.7 (θ = 36°).
The polarization resolved transients at 1682 cm−1 (Fig.3c)
show a dichroic ratio of about 1.6 (θ = 39°) at ~ 1ps delay
time, matching the expected value for the carbonyl groups
of the “capping helix”. The negative signals in the 12ps
DAS ranging from 1670 to 1600 cm−1 are assigned to apo-
protein dynamics of backbone and amino acid side chain
contributions. A more precise analysis is not possible here.
The negative signals at 1724 and 1708 cm−1 should origi-
nate from carboxylic or carbonyl C = O stretching vibrations
of amino acid side chains or from high-frequency arginine
(CN) stretching vibrations.
The constant DAS component in Fig.4b reflects changes
induced by the photoisomerization. Vibrations formed by
the photoproduct show positive features, while absent vibra-
tions from the parent state cause negative features. Posi-
tive/negative features at 1724 cm−1( +)/1705 cm−1(−) and
at 1680 cm−1( +)/1620 cm−1(−) are apparent. The peak pair
at 1724 cm−1( +)/1705 cm−1(−) was assigned to ring D car-
bonyl stretching vibration in the Lumi-F and ground state,
respectively. The intensity ratio of this peak pair was found
to be about 1:1 in labeled protein, in contrast to the pre-
sented ratio of 1:4 in unlabeled protein. We assign this dif-
ference to an additional long-living negative signal around
1724 cm−1 from an apoprotein group. The other peak pair
at 1680 cm−1( +)/1620 cm−1(−) was not observed in labeled
apoprotein. Thus, we assign this peak pair solely to apopro-
tein groups, in contrast to the assignment of the positive
1680 cm−1 contribution to a chromophore vibration [50].
Possible candidates for apoprotein vibrations are carbonyl
groups from the protein backbone, but other contributions
from amino acid side chains are also possible. Further stud-
ies have to be undertaken to disentangle individual apopro-
tein contributions.
In the spectral range between 1750 and 1660 cm−1 mainly
absorption bands from C = O stretching vibrations of the
apoprotein and PCB, and CN vibrations of arginine are
expected. Upon 13C/15N apoprotein labeling, the frequen-
cies of the C = O stretching vibrations downshift by about
40 cm−1, while a downshift of about 30 cm−1 is expected
for the high-frequency arginine (CN) stretching vibration
[50]. In 13C/15N labeled apoprotein the signal with the high-
est frequency is observed around 1725 cm−1. If the absorp-
tion difference signals between 1730 and 1690 cm−1 derived
from PCB alone, the absorbance difference spectra between
unlabeled and labeled protein would not be distinguishable
in this region. This is clearly not the case when comparing
unlabeled and labeled apoprotein in Fig.2a and 2c, respec-
tively. Hence, we observe apoprotein contributions in the
ultrafast dynamics prior to and after photoisomerization.
3.4 Apoprotein contributions
Direct comparison of the transients at 1724 cm−1 in unla-
beled and labeled apoprotein in Fig.3a indicate an instan-
taneous additional negative signal at this spectral position
in unlabeled apoprotein persisting up to 50ps after excita-
tion. An apoprotein bleaching band in the unlabeled apo-
protein around 1724 cm−1 should be visible as a red-shifted
bleaching band in the 13C/15N labeled apoprotein around
1680 cm−1. In Fig.4a the comparison between labeled (red
lines) and unlabeled (black lines) absorption difference
spectra are shown. At 50ps significant alterations between
labeled and unlabeled apoprotein are only visible around
1724 cm−1. The expected red-shifted negative bleaching
band around 1680 cm−1 is missing. It seems that this can
only be explained by the existence of another positive band
at the position of the expected red-shifted bleaching band
cancelling out the negative signal. Such a positive chromo-
phore band at 1680 cm−1 was reported and assigned to a
Lumi-F product band [50]. In the spectral range between
1660 and 1400 cm−1 various vibrational bands from the
protein backbone and side chains contribute to the overall
signal. Studies with different labeling strategies, e.g. with
and without chromophore labeling, and mutants will unravel
specific signal contributions.
Polarization resolved data show a dichroic ratio D < 1 of
the negative signal contribution at 1724 cm−1 after photo-
reaction. This corresponds to a relative angle in the range
of 54°–90° with respect to the electronic tdm. The negative
signal is generated instantaneously and persists on a time-
scale of 50ps. Such an apoprotein signal could indicate a
deprotonation of a carboxylic acid that stays deprotonated
after photoisomerization or a long lasting change of bond
character of a carbonyl group [34]. Possible candidates for
the signal are the carbonyl stretching vibration of the car-
boxylic acids of Asp207 and Glu196, as well as the carbonyl
stretching vibration of the backbone C = O group of Asp207.
All these groups are in the vicinity of the chromophore and
can easily be affected by the local electric field change of
the chromophore in the excited state. Moreover, their C = O
orientations exhibit relative angles in the range of 54°–90°
to the excited electronic tdm of the chromophore, consistent
with a dichroic ratio D < 1.
Up to now, no high-resolution structure of the Cph1
phytochrome in the Pfr state is available. Thus, conclusive
evidence on the protonation state of side groups, such as
Photochemical & Photobiological Sciences
1 3
glutamic and aspartic acids are scarce. Structural modelling
on the basis of solid-state magic-angle spinning NMR data
provide tentative structural information for the Pfr state in
Cph1 [12, 55]. Additional experimental data, such as polari-
zation resolved transient IR data, provide insight into hetero-
geneity of the ground state. On the basis of these data, two
ground-state conformations for the Pfr state were reported,
with deviating ring D orientations. The down-conformation
exhibits a ring D with an NH group pointing below the PCB
plane (see Fig.1c), while the up-conformation has a ring D
orientation with NH group pointing above the PCB plane
[20]. In these Pfr ground-state models possible protonation
of aspartic acids at pD 7.8 were not included. On the basis of
our observation of a bleaching band around 1724 cm−1, we
expect Asp207 to be protonated in the Pfr state. We changed
the protonation state of Asp207 in our model (see Fig.1c).
One CO group of Asp207 is hydrogen-bonded with the NH
group of PCB ring D with a distance of 1.85Å. This C = O
group is also hydrogen-bonded to Ser474 with a distance
of 1.57Å, and to Tyr263 with a distance of 2.89Å (see
Fig.1c). We assign the other CO group of Asp207 to be
protonated and hydrogen-bonded to Ser206 with a distance
of 2.02Å (see Fig.1c). The vtdm of the C = O group of
Asp207 shows a relative angle to the electronic tdm of 63°
compatible with the measured angle between 55 and 90°. Its
calculated frequency was at 1719 cm−1 with a relative inten-
sity of about 40% of the ring D carbonyl. These calculated
properties match with our experimental observations. The
vtdm of the Asp207 backbone carbonyl is oriented 79° to the
electronic tdm, also in agreement with the measured angle
between 55 and 90°. Its calculated frequency was found at
1728 cm−1 with a relative intensity of about 5% of the ring
D carbonyl. The small calculated relative intensity does not
match with experimental observations. In addition, the back-
bone carbonyl group would have to change its properties
completely, e.g. become protonated, to induce a long lasting
signal without an adjacent positive contribution. Another
possible candidate for the signal at 1724 cm−1 is Glu196,
since its carbonyl is also oriented roughly perpendicular to
the electronic tdm. However, its carboxylic group is about
12Å from the oxygen of ring D, making it rather unlikely
to be responsible for the instantaneous bleaching signal at
1724 cm−1.
There are a few groups that could be responsible for the
observed apoprotein dynamics at 1724 cm−1. We favor the
scenario, where a protonated Asp207 becomes deprotonated
upon photoexcitation.
The positive band around 1670 cm−1 in unlabeled apopro-
tein exhibits a dichroic ratio of about 1.8 pointing to a vtdm
with a relative angle of about 34° to the electronic tdm. The
spectral position matches frequencies of the α-helical amide
I vibration. The capping helix (see Fig.1) consists of many
amide I vibrations with relative angles of their C = O group
to the electronic tdm between 14° and 60°. Since the vtdm
deviates by about 10° from the C = O group direction of the
amide group, amide groups in the capping helix are suit-
able candidates for the observed signal around 1670 cm−1
in unlabeled apoprotein. Most of the other amide groups
in the phytochrome exhibit relative angles around 80° to
the electronic tdm, and thus do not match the observed sig-
nals. The short distance between the capping helix and the
chromophore supports such an assignment, since the cou-
pled dipoles of the amide I groups are affected more strongly
by an excited state dipole change the closer they are.
Negative signals around 1630 cm−1 are probably due to
apoprotein vibrations due to their substantial change upon
apoprotein labeling. Nevertheless, assignment is hampered
by increasing possibilities of overlapping signals. Further
deconvolution of the signal contributions can be performed
by including isotopic labeling of the PCB chromophore in
future experiments.
4 Discussion
4.1 Ultrafast protein dynamics
Our investigations support the notion that ultrafast photoi-
somerization of unlabeled PCB chromophore exhibit the
same dynamics and quantum yield in unlabeled and 13C/15N
labeled apoprotein. Comparison of 13C/15N labeled apopro-
tein and unlabeled apoprotein reveal ultrafast apoprotein
dynamics prior to and after photoisomerization of the PCB
chromophore, as revealed by isotope shifts.
At 1724 cm−1 a protein bleaching signal rises upon photo-
excitation. The bleaching signal remains almost constant on
a time-scale up to 50ps. We tentatively assign this band to
the C = O stretching vibration of Asp207 carboxylic acid that
is deprotonated upon photoisomerization. The deprotonated
carboxylic acid is expected to be relevant in down-stream
processes such as Arg-Asp salt bridge formation [56]. This
assignment is supported by polarization resolved data that
indicate a relative angle of the vtdm of the carbonyl group
to the electronic tdm of PCB between 55° and 90°, match-
ing the calculated angle of 63°. The impact of this aspartic
acid on the photoreaction of the Pr state was discussed for
bacteriophytochrome RpBphP3 from Rhodopseudomonas
palustris as a proton acceptor in the electronic excited state
[57]. In this report the backbone C = O group of Asp207 was
involved. This process is unlikely for the observed protein
signal at 1724 cm−1, because the signal remains after the
decay of the electronic excited state up to 50ps, but cannot
be excluded.
Excited state proton transfer from the chromophore to
the apoprotein was observed in the Pfr state of Agp2 phy-
tochrome in the electronic excited state. Its decay on a
Photochemical & Photobiological Sciences
1 3
sub-picosecond time-scale is accompanied by the re-proto-
nation of the chromophore [35]. A similar mechanism seems
very unlikely in Cph1 Pfr, however since no continuum band
is observed.
Alternative assignments of the 1724 cm−1 protein band
cannot be excluded. A possibility would be the arginine salt
bridge with the deprotonated propionic side chain at ring B.
The high frequency CN vibration of arginine in an Arg:Asp
salt bridge was simulated to absorb around 20 cm−1 red-
shifted to the C = O stretching vibration of ring D [50]. This
is in line with other studies assigning arginine salt bridges
at lower frequencies around 1620 cm−1 [58]. Hence, the
observed apoprotein band exhibits a frequency probably too
high for arginine salt bridges. Other glutamic and aspar-
tic acids, perhaps Glu196 could match the frequency and
the measured orientation. Since the response of the signal
is concomitant with PCB photoisomerization, we expect a
direct coupling to the chromophore or a coupling via hydro-
gen bonds. This is obvious for Asp207, but not clear for
other carboxylic acids in the protein.
The positive signal around 1670 cm−1 in unlabeled pro-
tein appears with photoexcitation of the Pfr state and decays
with a time constant of about 12ps to a small positive value
at 50ps. Its vtdm exhibits a relative angle to the electronic
tdm of about 35°. The underlying chromophore contribution
is negligible. This relative angle agrees with many backbone
carbonyl groups of the capping helix that is oriented parallel
to electronic tdm of PCB (see Fig.1b). This corresponds to
ultrafast structural changes observed in the capping helix by
serial femtosecond X-ray crystallography in the Pr state of
DrBphP from Deinococcus radiodurans [37].
4.2 Proposed mechanism
Upon excitation of Pfr, the electron density is increased
around ring B and decreased around rings C and D of the
chromophore [35]. This almost instantaneous event induces
an impulsive electric field change around the chromophore.
This process has been called a ‘Coulomb Hammer’, alter-
ing the coupling strengths of hydrogen bonds between the
chromophore and the surrounding protein [36]. Polarizable
and charged protein groups will be directly affected by the
electric field change, before reorientation of movable groups,
e.g. water, accompanied with screening of the electric field
takes place. The effects on individual groups can vary con-
siderably depending on polarizability and are currently
difficult to quantify. Further studies are needed to unravel
transient effects on individual groups.
We assign the apoprotein bleaching band at 1724 cm−1
tentatively to the deprotonation of Asp207. With excitation
and the instantaneous decrease of electron density at ring D
and increase around ring B the hydrogen-bonding network
between the NH group of ring D, Asp207, Ser474, and
Tyr263 is altered. We propose a weakening of the hydrogen
bond to ring D, facilitating its rotation. This is necessary,
because strong hydrogen bonds can impede photoisomeriza-
tion—as demonstrated for bacteriophytochrome RpBphP2
and RpBphP3 from Rhodopseudomonas palustris [59]. With
the weakening of the hydrogen bond to ring D and beginning
rotation of ring D, the hydrogen-bonding network rearranges
and Asp207 is deprotonated. Up to now, we were not able
to identify the proton acceptor. These investigations can be
carried out in a more targeted manner once a high-resolution
structure of the Cph1 Pfr state becomes available.
Ultrafast apoprotein responses around 1680 cm−1 were
assigned to amide I vibrations. We assigned the interacting
amide I groups to the capping helix above and parallel to the
chromophore. The electric field change of the chromophore
polarizes the α-helix and strengthens the couplings of the
amide I groups, thereby increasing their absorption result-
ing in the observed positive signal around 1670 cm−1 [23].
By comparing unlabeled with labeled apoprotein dynam-
ics we were able to identify ultrafast apoprotein responses
prior to and after the photoisomerization of the PCB
chromophore in Cph1. This highlights the impact of the
excitation induced electric field change on the apoprotein
response and supports the ‘Coulomb hammer’ mechanism
setting the protein up for photoisomerization of the chromo-
phore and down-stream protein changes.
Acknowledgements The work was funded by the Deutsche Forschun-
gsgemeinschaft (DFG) through project-ID 221545957−CRC 1078
‘Protonation Dynamics in Protein Function’, subproject B07 (K.H.,
and J.H.), subprojects C02 (M.A.M.) and through DFG grant project
HE 5206/3-2 (K.H.) and HU 702/12-1 (J.H.).
Funding Open Access funding enabled and organized by Projekt
DEAL.
Data availability The datasets generated during and/or analysed dur-
ing the current study are available from the corresponding author on
reasonable request.
Declarations
Conflict of interest The authors have no competing interests to declare
that are relevant to the content of this article.
Open Access This article is licensed under a Creative Commons Attri-
bution 4.0 International License, which permits use, sharing, adapta-
tion, distribution and reproduction in any medium or format, as long
as you give appropriate credit to the original author(s) and the source,
provide a link to the Creative Commons licence, and indicate if changes
were made. The images or other third party material in this article are
included in the article's Creative Commons licence, unless indicated
otherwise in a credit line to the material. If material is not included in
the article's Creative Commons licence and your intended use is not
permitted by statutory regulation or exceeds the permitted use, you will
need to obtain permission directly from the copyright holder. To view a
copy of this licence, visit http:// creat iveco mmons. org/ licen ses/ by/4. 0/.
Photochemical & Photobiological Sciences
1 3
References
1. Heyne, K., etal. (2002). Ultrafast dynamics of phytochrome
from the cyanobacterium Synechocystis, reconstituted with phy-
cocyanobilin and phycoerythrobilin. Biophysical Journal, 82,
1004–1016.
2. Rohmer, T., etal. (2010). Phytochrome as molecular machine:
revealing chromophore action during the Pfr -> Pr photoconver-
sion by magic-angle spinning NMR spectroscopy. Journal of the
American Chemical Society, 132, 9219–9219. https:// doi. org/ 10.
1021/ Ja104 192u
3. Foerstendorf, H., Lamparter, T., Hughes, J., Gartner, W., & Sie-
bert, F. (2000). The photoreactions of recombinant phytochrome
from the cyanobacterium Synechocystis: a low-temperature UV-
Vis and FT-IR spectroscopic study. Photochemistry and Photobi-
ology, 71, 655–661.
4. Foerstendorf, H., etal. (2001). FTIR studies of phytochrome
photoreactions reveal the C=O bands of the chromophore: Con-
sequences for its protonation states, conformation, and protein
interaction. Biochemistry, 40, 14952–14959.
5. Kim, P. W., Rockwell, N. C., Martin, S. S., Lagarias, J. C., &
Larsen, D. S. (2014). Heterogeneous photodynamics of the P-fr
State in the Cyanobacteriai phytochrome Cph1. Biochemistry, 53,
4601–4611.
6. Muller, M. G., Lindner, I., Martin, I., Gartner, W., & Holzwarth,
A. R. (2008). Femtosecond kinetics of photoconversion of the
higher plant photoreceptor phytochrome carrying native and
modified chromophores. Biophysical Journal, 94, 4370–4382.
7. Schumann, C., etal. (2008). Subpicosecond midinfrared spectros-
copy of the P-fr reaction of phytochrome Agp1 from Agrobacte-
rium tumefaciens. Biophysical Journal, 94, 3189–3197. https://
doi. org/ 10. 1529/ bioph ysj. 107. 119297
8. Wang, D., etal. (2019). Elucidating the molecular mechanism
of ultrafast Pfr state photoisomerization in Bathy Bacteriophy-
tochrome PaBphP. Journal of Physical Chemistry Letters, 10,
6197–6201.
9. Escobar, F. V., etal. (2015). Conformational heterogeneity of the
Pfr chromophore in plant and cyanobacterial phytochromes. Fron-
tiers in Molecular Biosciences. https:// doi. org/ 10. 3389/ fmolb.
2015. 00037
10. Fischer, T., etal. (2022). Influence of the PHY domain on the
ms-photoconversion dynamics of a knotless phytochrome. Pho-
tochemical and Photobiological Sciences. https:// doi. org/ 10. 1007/
s43630- 022- 00245-9
11. Fischer, T., etal. (2021). Ultrafast photoconversion dynamics
of the knotless phytochrome SynCph2. International Journal of
Molecular Sciences. https:// doi. org/ 10. 3390/ ijms2 21910 690
12. Song, C., etal. (2018). 3D Structures of plant phytochrome A
as Pr and Pfr from solid-state NMR: implications for molecular
function. Frontiers in Plant Science. https:// doi. org/ 10. 3389/ fpls.
2018. 00498
13. Legris, M., Ince, Y. C., & Fankhauser, C. (2019). Molecular
mechanisms underlying phytochrome-controlled morphogen-
esis in plants. Nature Communications. https:// doi. org/ 10. 1038/
s41467- 019- 13045-0
14. Anders, K., Gutt, A., Gartner, W., & Essen, L. O. (2014). Photo-
transformation of the red light sensor cyanobacterial Phytochrome
2 from Synechocystis species depends on its tongue motifs. Jour-
nal of Biological Chemistry, 289, 25590–25600.
15. Bischoff, M., Hermann, G., Rentsch, S., & Strehlow, D. (2001).
First steps in the phytochrome phototransformation: a comparative
femtosecond study on the forward (Pr -> Pfr) and back reaction
(Pfr -> Pr). Biochemistry, 40, 181–186.
16. Essen, L. O., Mailliet, J., & Hughes, J. (2008). The structure of
a complete phytochrome sensory module in the Pr ground state.
Proceedings of the National Academy of Sciences USA, 105,
14709–14714. https:// doi. org/ 10. 1073/ pnas. 08064 77105
17. Nagano, S., etal. (2020). Structural insights into photoactivation
and signalling in plant phytochromes. Nature Plants. https:// doi.
org/ 10. 1038/ s41477- 020- 0638-y
18. Takala, H., etal. (2014). Signal amplification and transduction in
phytochrome photosensors. Nature, 509, 245–248.
19. Yang, X., Kuk, J., & Moffat, K. (2008). Crystal structure of Pseu-
domonas aeruginosa bacteriophytochrome: photoconversion and
signal transduction. Proceedings of the National Academy of
Sciences USA, 105, 14715–14720. https:// doi. org/ 10. 1073/ pnas.
08067 18105
20. Stensitzki, T., etal. (2017). Influence of heterogeneity on the ultra-
fast photoisomerization dynamics of Pfr in Cph1 phytochrome.
Photochemistry and Photobiology, 93, 703–712.
21. Yang, Y., Heyne, K., Mathies, R. A., & Dasgupta, J. (2016). Non-
bonded interactions drive the sub-picosecond bilin photoisomeri-
zation in the P-fr state of phytochrome Cph1. ChemPhysChem, 17,
369–374.
22. Inomata, K., etal. (2005). Sterically locked synthetic bilin
derivatives and phytochrome Agp1 from Agrobacterium tume-
faciens form photoinsensitive Pr- and Pfr-like adducts. Journal
of Biological Chemistry, 280, 24491–24497.
23. Gross, R., etal. (2009). Ultrafast protein conformational alter-
ations in bacteriorhodopsin and its locked analogue BR5.12.
Journal of Physical Chemistry B, 113, 7851–7860.
24. Linke, M., etal. (2013). Electronic transitions and heterogeneity
of the bacteriophytochrome Pr absorption band: An angle bal-
anced polarization resolved femtosecond VIS pump-IR probe
study. Biophysical Journal, 105, 1756–1766.
25. Fan, G. B., Siebert, F., Sheves, M., & Vogel, R. (2002). Rho-
dopsin with 11-cis-locked chromophore is capable of forming
an active state photoproduct. Journal of Biological Chemistry,
277, 40229–40234.
26. Zhong, Q., etal. (1996). Reexamining the primary light-induced
events in bacteriorhodopsin using a synthetic C-13=C-14-
locked chromophore. Journal of the American Chemical Soci-
ety, 118, 12828–12829.
27. Foster, K. W., Saranak, J., Krane, S., Johnson, R. L., & Nakani-
shi, K. (2011). Evidence from Chlamydomonas on the photoac-
tivation of rhodopsins without isomerization of their chromo-
phore. Chemistry and Biology, 18, 733–742. https:// doi. or g/ 10.
1016/j. chemb iol. 2011. 04. 009
28. Salem, L., & Bruckmann, P. (1975). Conversion of a photon to
an electrical signal by sudden polarisation in the N-retinylidene
visual chromophore. Nature, 258, 526–528. https:// doi. org/ 10.
1038/ 25852 6a0
29. Mathies, R., & Stryer, L. (1976). Retinal has a highly dipolar
vertically excited singlet state: implications for vision. Proceed-
ings of the National Academy of Sciences USA, 73, 2169–2173.
https:// doi. org/ 10. 1073/ pnas. 73.7. 2169
30. Lewis, A. (1978). The molecular mechanism of excitation in
visual transduction and bacteriorhodopsin. Proceedings of the
National Academy of Sciences USA, 75, 549–553. https:// doi.
org/ 10. 1073/ pnas. 75.2. 549
31. Neumann-Verhoefen, M. K., etal. (2013). Ultrafast infrared
spectroscopy on channel rhodopsin-2 reveals efficient energy
transfer from the retinal chromophore to the protein. Journal of
the American Chemical Society, 135, 6968–6976. https:// doi.
org/ 10. 1021/ ja400 554y
32. Das, I., Pushkarev, A., & Sheves, M. (2021). Light-induced
conformational alterations in heliorhodopsin triggered by the
retinal excited state. The Journal of Physical Chemistry B, 125,
8797–8804. https:// doi. org/ 10. 1021/ acs. jpcb. 1c045 51
33. Tahara, S., Kuramochi, H., Takeuchi, S., & Tahara, T. (2019).
Protein dynamics preceding photoisomerization of the retinal
Photochemical & Photobiological Sciences
1 3
chromophore in bacteriorhodopsin revealed by deep-UV fem-
tosecond stimulated Raman spectroscopy. Journal of Physical
Chemistry Letters, 10, 5422–5427.
34. Stensitzki, T., Adam, S., Schlesinger, R., Schapiro, I., & Heyne,
K. (2020). Ultrafast backbone protonation in channel rhodop-
sin-1 captured by polarization resolved Fs Vis-pump-IR-Probe
spectroscopy and computational methods. Molecules. https://
doi. org/ 10. 3390/ molec ules2 50408 48
35. Yang, Y., etal. (2022). Ultrafast proton-coupled isomerization
in the phototransformation of phytochrome. Nature Chemistry,
14, 823–830.
36. Heyne, K. (2022). Impact of ultrafast electric field changes on
photoreceptor protein dynamics. The Journal of Physical Chem-
istry B, 126, 581–587.
37. Claesson, E., etal. (2020). The primary structural photore-
sponse of phytochrome proteins captured by a femtosecond
X-ray laser. eLife. https:// doi. org/ 10. 7554/ eLife. 53514
38. Nogly, P., etal. (2018). Retinal isomerization in bacteriorhodopsin
captured by a femtosecond x-ray laser. Science. https:// doi. org/ 10.
1126/ scien ce. aat00 94
39. Kovacs, G. N., etal. (2019). Three-dimensional view of ultrafast
dynamics in photoexcited bacteriorhodopsin. Nature Communica-
tions. https:// doi. org/ 10. 1038/ s41467- 019- 10758-0
40. Yang, Y., etal. (2012). Real-time tracking of phytochrome’s ori-
entational changes during Pr photoisomerization. Journal of the
American Chemical Society, 134, 1408–1411.
41. Yang, Y., etal. (2014). Active and silent chromophore isoforms
for phytochrome Pr photoisomerization: an alternative evolution-
ary strategy to optimize photoreaction quantum yields. Structural
Dynamics. https:// doi. org/ 10. 1063/1. 48652 33
42. Hahn, J., Strauss, H. M., & Schmieder, P. (2008). Heteronuclear
NMR investigation on the structure and dynamics of the chromo-
phore binding pocket of the cyanobacterial phytochrome Cph1.
Journal of the American Chemical Society, 130, 11170–11178.
https:// doi. org/ 10. 1021/ Ja803 1086
43. Roben, M., etal. (2010). NMR Spectroscopic investigation of
mobility and hydrogen bonding of the chromophore in the binding
pocket of phytochrome proteins. ChemPhysChem, 11, 1248–1257.
https:// doi. org/ 10. 1002/ cphc. 20090 0897
44. Schmieder, P., Strauss, H. M., & Hughes, J. (2005). Heteronuclear
solution-state NMR studies of the chromophore in cyanobacterial
phytochrome Cph1. Biochemistry, 44, 8244–8250. https:// doi. or g/
10. 1021/ Bi050 457r
45. Song, C., etal. (2011). Two ground state isoforms and a chromo-
phore D-ring photoflip triggering extensive intramolecular
changes in a canonical phytochrome. Proceedings of the National
Academy of Sciences USA, 108, 3842–3847.
46. Mroginski, M. A., Mark, F., Thiel, W., & Hildebrandt, P. (2007).
Quantum mechanics/molecular mechanics calculation of the
Raman spectra of the phycocyanobilin chromophore in alpha-c-
phycocyanin. Biophysical Journal, 93, 1885–1894.
47. Mroginski, M. A., etal. (2011). Elucidating photoinduced struc-
tural changes in phytochromes by the combined application of
resonance Raman spectroscopy and theoretical methods. Journal
of Molecular Structure, 993, 15–25.
48. Gaussian09 (2009).
49. Levina, E. O., Lokshin, B. V., Mai, B. D., & Vener, M. V. (2016).
Spectral features of guanidinium-carboxylate salt bridges. The
combined ATR-IR and theoretical studies of aqueous solution of
guanidinium acetate. Chemical Physics Letters, 659, 117–120.
50. Ihalainen, J. A., etal. (2018). Chromophore-protein interplay
during the phytochrome photocycle revealed by step-scan FTIR
spectroscopy. Journal of the American Chemical Society, 140,
12396–12404.
51. Braiman, M. S., Briercheck, D. M., & Kriger, K. M. (1999). Mod-
eling vibrational spectra of amino acid side chains in proteins:
effects of protonation state, counterion, and solvent on arginine
C-N stretch frequencies. The Journal of Physical Chemistry B,
103, 4744–4750.
52. Lotze, S., & Bakker, H. J. (2015). Structure and dynamics of a
salt-bridge model system in water and DMSO. The Journal of
Chemical Physics. https:// doi. org/ 10. 1063/1. 49189 04
53. Escobar, F. V., etal. (2015). A protonation-coupled feedback
mechanism controls the signalling process in bathy phytochromes.
Nature Chemistry, 7, 423–430.
54. Theisen, M., etal. (2009). Femtosecond polarization resolved
spectroscopy: A tool for determination of the three-dimensional
orientation of electronic transition dipole moments and identifica-
tion of configurational isomers. The Journal of Chemical Physics.
https:// doi. org/ 10. 1063/1. 32368 04
55. Song, C., etal. (2013). Solid-state NMR spectroscopy to probe
photoactivation in canonical phytochromes. Photochemistry and
Photobiology, 89, 259–273. https:// doi. org/ 10. 1111/ php. 12029
56. Mroginski, M. A., etal. (2009). Chromophore structure of cyano-
bacterial phytochrome Cph1 in the Pr state: reconciling structural
and spectroscopic data by QM/MM calculations. Biophysical
Journal, 96, 4153–4163.
57. Toh, K. C., Stojkovic, E. A., van Stokkum, I. H. M., Moffat, K.,
& Kennis, J. T. M. (2010). Proton-transfer and hydrogen-bond
interactions determine fluorescence quantum yield and photo-
chemical efficiency of bacteriophytochrome. Proceedings of the
National academy of Sciences of the United States of America,
107, 9170–9175.
58. Huerta-Viga, A., etal. (2015). The structure of salt bridges
between Arg(+) and Glu(-) in peptides investigated with 2D-IR
spectroscopy: Evidence for two distinct hydrogen-bond geome-
tries. The Journal of Chemical Physics. https:// doi. org/ 10. 1063/1.
49210 64
59. Toh, K. C., etal. (2011). Primary Reactions of bacteriophy-
tochrome observed with ultrafast mid-infrared spectroscopy.
Journal of Physical Chemistry A, 115, 11985–11997.
