Regular Paper

3D Confocal Laser Scanning Microscopy for the Analysis of
Chlorophyll Fluorescence Parameters of Chloroplasts in Intact
Leaf Tissues
Kenji Omasa*, Atsumi Konishi, Hikaru Tamura and Fumiki Hosoi
Graduate School of Agricultural and Life Sciences, The University of Tokyo, 1-1-1 Yayoi, Bunkyo, Tokyo, 113-8675 JapanWe analyzed the chlorophyll ﬂuorescence parameters in
a 3D cellular arrangement in vivo by using a modiﬁed
Nipkow disk-type confocal laser scanning microscope (CLSM).
We ﬁrst deﬁned the 3D values of ΦPSII (photochemicalyield of PSII) and NPQ (non-photochemical quenching) inmesophyll, epidermal and guard cell chloroplasts from theleaf surface to several tens of microns in depth. We also used
this CLSM method to analyze the relationships between
actinic light intensity and the chlorophyll ﬂuorescenceparameters for Boston fern and broad bean leaf specimens.
As the actinic light intensity increased, the mean ΦPSII
values decreased and the NPQ values increased in all
chloroplasts of Boston fern and broad bean leaf. These
values differed with cell type and species. The Boston fern
chloroplasts had lower ΦPSII values than the broad beanchloroplasts, and vice versa for the NPQ values. The ΦPSII
values of Boston fern chloroplasts decreased in the ordermesophyll, epidermal and guard cell chloroplasts. The
NPQ values decreased in the order guard cell, mesophylland epidermal chloroplasts, except at 12 µmol m–2 s–1
actinic light, when the mesophyll value was slightly lower
than that of the epidermis. The trend in the ΦPSII and NPQ
values of broad bean mesophyll and guard cell chloroplasts
was opposite to that of Boston fern chloroplasts. As 3D
CLSM can provide the ΦPSII and NPQ values of eachchloroplast in a 3D cellular arrangement, this method has
potential for investigating differences in the functions ofchloroplasts in vivo.
Keywords: 3D • Chlorophyll ﬂuorescence • Chloroplast •CLSM • NPQ • ΦPSII.
Abbreviations: 3D, three dimensional; a, absorption coefﬁcientof photosynthetic pigments; A/D, analog to digital; CLSM,
confocal laser scanning microscopy; iDt, dark current
image integrated for t ms; EM-CCD, electron-multiplying

charge-coupled device; ETR, electron transport rate; iF,
chlorophyll ﬂuorescence intensity image measured underactinic light; iFm, chlorophyll ﬂuorescence intensity image
measured during a saturation light pulse in darkness; iFm′,
chlorophyll ﬂuorescence intensity image measured during
the saturation light pulse under actinic light; I, irradiated lightintensity; ND, neutral density; NPQ, non-photochemical
quenching of chlorophyll ﬂuorescence; PPF, photosynthetic
photon ﬂux; QA, primary quinone acceptor of PSII; QB, secondary
quinone acceptor of PSII; R2, coefﬁcient of determination;
ΦF, chlorophyll ﬂuorescence yield under actinic light; ΦFm,
chlorophyll ﬂuorescence yield during a saturation light pulsein the dark; ΦFm′, chlorophyll ﬂuorescence yield during asaturation light pulse under actinic light; ΦF0, minimum
chlorophyll ﬂuorescence yield after dark adaptation; ΦFv,
ΦFm–ΦF0,; ΦFv/ΦFm, maximum photochemical yield of PSII
after dark adaptation; ΦPSII, photochemical yield of PSII; λ,wavelength of light.

Introduction
Chlorophyll ﬂuorescence imaging is one of the leading methods used to assess plant photosynthetic activities, because
information is provided without the need for destruction of,or contact with, the living leaves (Omasa et al. 1987, Daleyet al. 1989, Genty and Meyer 1995, Rolfe and Scholes 1995,
Lichtenthaler and Miehe 1997, Govindjee and Nedbal 2000,
Omasa and Takayama 2002, Osmond and Park 2002, Omasa
and Takayama 2003, Papageorgiou and Govindjee 2004,
Chaerle et al. 2007, Omasa et al. 2007). Chlorophyll ﬂuorescence microscopy is an advanced imaging technique used todetect photosynthetic activity at the level of individual cells
and chloroplasts in a non-invasive manner (Oxborough2004). The ﬁrst chlorophyll ﬂuorescence microscopy technique was developed by Oxborough and Baker (1997a) andapplied the ‘saturation pulse method’ for calculating ΦPSII,*Corresponding author: Email, aomasa@mail.ecc.u-tokyo.ac.jp; Fax,+81-3-5841-8175.

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174, available online at www.pcp.oxfordjournals.org© The Author 2008. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
All rights reserved. For permissions, please email: journals.permissions@oxfordjournals.org

90

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

3D CLSM of chlorophyll ﬂuorescence parameterswhich represents the quantum yield of photosynthetic
electron transport (Genty et al. 1989). This technique has since
been used for applications that include comparing responses
at the chloroplast level in mesophyll and guard cells with changes
in ambient CO2, O2, light and humidity levels (Oxboroughand Baker 1997a, Oxborough and Baker 1997b, Baker et al.
2001, Lawson et al. 2002, Lawson et al. 2003); investigating
the effects of exposing leaf cells to ozone (Leipner et al. 2001);
assessing the heterogeneity of cells and coenobia of microalgae (Küpper et al. 2000); and detecting herbicide stress inalgae at different growth stages (Endo and Omasa 2004).
The generation of 3D information on the inherently complicated 3D structures of leaf tissues, cells and chloroplasts isrequired to improve our understanding of plant functioning
and responses to stresses (Rigaut et al. 1992, Omasa 2000,
Schurr et al. 2006, Omasa et al. 2007). In 3D surface microscopy of chlorophyll ﬂuorescence, extended-focus ΦPSII imagesof the leaf tissue surface were reconstructed from 39 imagesof chlorophyll ﬂuorescence, taken at different focal planes at
5.5 µm intervals and with a 10× objective lens (Rolfe andScholes 2002). Thereafter, the reconstruction of 3D cell-level
ΦPSII from <15 images taken at 4 µm intervals was achievedby using a modiﬁed shape-from-focus algorithm and higher
magniﬁcation (×20) lens (Endo and Omasa 2007). Despite
the highly efﬁcient methods for reconstruction used by thesetwo previous studies, the internal 3D anatomy of tissues andcells was not visualized, as the 3D surface reconstruction was
performed passively. Furthermore, the objective magniﬁcations were not sufﬁcient to capture high-resolution imagesof chloroplasts within the cells.
Confocal laser scanning microscopy (CLSM), invented by
Minsky (1988), is an active microscopic technique with thecapacity for higher resolution and performance for 3D
reconstruction than conventional passive light microscopy(Rigaut et al. 1992). This is because CLSM eliminates outof-focus haze in objects by passing the excitation laser beamand the reﬂected or ﬂuorescent light from the objects
through pinholes (Nakano 2002). CLSM can also reconstruct
images of the internal 3D anatomy of tissues and cells by
capturing the ﬂuorescence from ﬂuorochromes within thetissues and cells. These advantages have enabled the use of
CLSM for chlorophyll ﬂuorescence microscopy of leaf tissue
proﬁles (Osmond et al. 1999) and of chloroplast movementin vivo (Tlałka and Fricker 1999, Tlałka et al. 1999). One disadvantage of the galvanometer-mirror-type CLSM approachis the requirement for specimens to be irradiated with astrong excitation laser beam in order to capture clear images;
the use of this galvanometer-mirror makes it technically difﬁcult to scan and evenly irradiate specimens rapidly (Wanget al. 2005). These issues prevent the use of the saturation
pulse method with CLSM for obtaining the ΦPSII and NPQ
chlorophyll ﬂuorescence parameters. Recent advances inhigh-speed rotating pinhole disk (Nipkow disk) CLSM

(Ichihara et al. 1996, Tanaami et al. 2002) and highly sensitive
cooled electron-multiplying charge-coupled device (EM-CCD)
technologies have made it possible to scan rapidly, evenly
irradiate and capture clear internal ﬂuorescence images of a
specimen under a low-intensity laser beam that has approximately 1/1,000 the intensity of the galvanometer-mirror-typelaser beam. The large number of confocal pinholes in thehigh-speed rotating disk makes it possible to use this lower
intensity beam. Recently, we developed a novel computeraided CLSM system for applying the saturation pulse method
through a modiﬁcation of the Nipkow disk CLSM andthe use of a high sensitivity EM-CCD camera (Omasa andKonishi 2008). This system can be used to analyze theinternal 3D anatomy of leaf tissues and cells and to assess
differences in chlorophyll ﬂuorescence parameters in vivo
for each chloroplast at the ordinary range of actinic lightintensity, using objectives >40×. In our previous study we
did not apply the system to the estimation of ΦPSII and NPQ
chlorophyll ﬂuorescence parameters, so the performance ofthe system was not assessed. For this reason, in our current
study, we investigated the performance of this system andassessed differences in the chlorophyll ﬂuorescence parameters of epidermal, guard cell and mesophyll chloroplasts inreconstructed 3D images of Boston fern (Nephrolepis exaltata L. ‘Bostoniensis’) and broad bean (Vicia faba L. ‘Komasakae’) leaf specimens under different actinic light intensities.

Results
Conﬁrmation of saturation light pulse intensity andduration of saturation of photosynthetic electrontransport
We examined the relationships between the intensity [photosynthetic photon ﬂux (PPF)] of the light pulse falling on
the Boston fern leaf and the chlorophyll ﬂuorescence yieldsof mesophyll and guard cell chloroplasts, as measured fromthe abaxial leaf surface by the CLSM system, and the tissuelevel yields at the abaxial leaf surface, as measured witha conventional chlorophyll ﬂuorescence imaging system(see Materials and Methods, Omasa and Takayama 2003)(Fig. 1). Fig. 1A and B shows the results for a dark-adaptedleaf and a leaf adapted under light of 300 µmol m–2 s–1 PPF,
respectively. The chlorophyll ﬂuorescence yield (ratio) on thevertical axis is the ratio of chlorophyll ﬂuorescence intensity
at each PPF to the average saturated intesnity. The ﬂuorescence yields of the dark-adapted leaf increased dramaticallywith a slight increase in light intensity, reaching approximately
1.0 at a PPF of 800 µmol m–2 s–1 (dotted line in Fig. 1A). The
ﬂuorescence yields of the light-adapted leaf increased more
gradually than those of the dark-adapted leaf, reachingapproximately 1.0 at a PPF of 1,300 µmol m–2 s–1 (dotted line
in Fig. 1B). The same trends were observed in the changes in

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

91

K. Omasa et al.

A

B

Chl. fluorescence yield (ratio)

1.0

1.0

0.8

0.8

800 mmol m−2 s−1

0.6

0.6

0.4

0.4

0.2

0.2

0.0

0

1000

2000

0.0

3000

1300 mmol m−2 s−1

0

1000

PPF (mmol m−2 s−1)

2000

3000

PPF (mmol m−2 s−1)

Fig. 1 Relationships between light pulse intensity (PPF) and chlorophyll ﬂuorescence yields of mesophyll and guard cell chloroplasts of Boston
fern leaf specimens (as measured from the abaxial leaf side with the CLSM system) and chlorophyll ﬂuorescence yield of the abaxial leaf surface(as measured with a conventional chlorophyll ﬂuorescence imaging system). (A) Results in the dark. (B) Results in 300 µmol m–2 s–1 actinic light.
Open squares, circles and triangles represent chlorophyll ﬂuorescence yields of mesophyll chloroplasts, guard cell chloroplasts and the abaxial
leaf surface, respectively. The yield values were the averages of 20 chloroplasts for mesophyll cells and 10 chloroplasts for guard cell chloroplasts.
Dotted lines represent the minimal PPF for the saturation light pulse.

Reconstruction of 3D chlorophyll ﬂuorescenceintensity images and calculation of chlorophyll
ﬂuorescence parameters
We examined three chlorophyll ﬂuorescence intensity imagesof a Boston fern leaf specimen selected from a series of images

92

2.9 s

0.12 s1.0
Chl. fluorescence intensity (ratio)

the ﬂuorescence yield of mesophyll and guard cell chloroplasts, as measured with the CLSM system, and in the tissuelevel yields of the abaxial leaf surface, as measured with theconventional imaging system.
We measured the chlorophyll ﬂuorescence transients of
Boston fern mesophyll chloroplasts from the abaxial leaf sidewith the CLSM system under different saturation light intensities(Fig. 2). Fig.2a, b and c shows the results for leaves acclimatized to actinic light at 248, 98 and 0 µmol m–2 s–1, respectively. The saturation light intensity was 4,820 µmol m–2 s–1 in
0 µmol m–2 s–1 actinic light and 24.33 times the actinic lightintensity in 248 and 98 µmol m–2 s–1 actinic light. The arrows
indicate the duration of saturation, at >97% of the maximum intensity, in each transient. The ﬂuorescence intensity
increased rapidly after irradiation under the saturation lightintensity and reached 97% of the maximum intensity in
0.12 s under actinic light at 248 and 98 µmol m–2 s–1 and in
0.15 s under actinic light at 0 µmol m–2 s–1. The duration ofsaturation ranged from 2.2 to 2.9 s, with the largest duration
associated with the highest intensity of actinic light. We
reconstructed a 3D image by capturing a series of chlorophyll ﬂuorescence intensity images during the saturation
laser light pulse in the dark and light in 64 different focal
planes for 1.92 s from 0.2 s after the start of irradiation.

a

0
1.0

2.8 s

0.12 s

b
0
1.0

2.2 s

0.15 s

c

0

0.01

0.1

1

10

100

Time (s)Fig. 2 Chlorophyll ﬂuorescence transients of mesophyll chloroplastsof Boston fern leaf specimens, as measured from the abaxial leaf sidewith the CLSM system under different saturation light intensities. The
chlorophyll fluorescence intensity was the average of 20 mesophyll
chloroplasts. a, b and c represent the results for leaves acclimatized
to 248, 98 and 0 µ mol m –2 s –1 actinic light PPF, respectively. The
saturation light was 4,820 µmol m–2 s–1 PPF in 0 µmol m–2 s–1 actinic
light and 24.33 times the actinic light in 248 and 98 µmol m–2 s–1
actinic light PPF. Durations from the peak intensities to 97% of thepeak intensities are indicated by arrows wedged between dotted lines.

captured in 64 different focal planes during irradiation for
approximately 2 s with a saturation light pulse of 4,820µmol m–2 s–1 PPF (Fig. 3A). We also examined 3D images

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

3D CLSM of chlorophyll ﬂuorescence parameters

reconstructed from the 64 planes before 3D deconvolution(Fig. 3B), and the images after 3D deconvolution (Fig. 3C).
Fig. 3D is a transmitted light image of the same microscopic
abaxial leaf area. In samples of A(i), i = 10, 30 and 50, where ‘i’is the order of focal planes with different z coordinates atincreasing depth from the abaxial leaf surface to the inside ofthe leaf. As the z-scan interval was 1.25 µm, A(10), A(30) andA(50) were at focal plane depths of 12.5, 37.5 and 62.5 µm,
respectively, from the leaf surface. The distance from the leaf
surface to A(64) was 80 µm. Focused and defocused ﬂuorescence areas are apparent in Fig. 3A. In A(10), ﬂuorescentchloroplasts in guard cells appear at the center of the image.
Some brightly ﬂuorescent chloroplasts in the epidermal cells
are also apparent. In A(30), the guard cell chloroplasts are
not apparent and a large number of ﬂuorescent mesophyll
chloroplasts are found, especially along the boundaries of
the cells. In A(50), a number of ﬂuorescent chloroplasts arefound on the right side of the focal plane. Spike noises in all
focal planes were ﬁrst eliminated by applying a median ﬁlter
with a mask size of 3 × 3 pixels.
The 3D chlorophyll ﬂuorescence intensity images werereconstructed by building the acquired planes. Fig. 3B is areconstructed 3D image from the abaxial surface and theside. The conﬁguration of the ﬂuorescent chloroplasts in thereconstructed 3D images reﬂected the conﬁguration normally observed in vivo. The reconstructed images were thendeconvoluted to obtain a clearer conﬁguration (Fig. 3C).
The deconvolution image was clearer, but the analog to digital (A/D) conversion level of the chlorophyll ﬂuorescenceintensity was inaccurate. Therefore, we used the reconstructed images before the deconvolution to calculate theΦPSII and NPQ values, and used the deconvolution image to
produce a 1-bit binary image to extract only the chloroplast
images (see Materials and Methods). Fig. 3D is a transmitted

Chl. fluo. int. (A/D conv.)
12000
2000

Chl. fluo. int. (A/D conv.)
2000
12000

Chl. fluo. int. (A/D conv.)
2000
12000

A (10)A (30)A (50)

Chl. fluo. int. (A/D conv.)
12000
2000

Chl. fluo. int. (A/D conv.)
216
0

B

C

D

30 mm

30 mm

Fig. 3 (A) Chlorophyll ﬂuorescence intensity images of Boston fern leaf specimen, selected from a series of images captured at 64 different
focal planes (= different z coordinates) during irradiation for approximately 2 s with a saturation light pulse. The value in parentheses for each
image in A (10, 30, 50) is the number of the focal plane from the abaxial leaf surface to the inside of the leaf. The saturation light pulse was
4,820 µmol m–2 s–1 PPF. (B) 3D images reconstructed from the 64 planes before 3D deconvolution. (C) 3D deconvoluted images. (D) Transmitted
light image of the same microscopic abaxial leaf area. The images from the surface and proﬁle are shown in B and C.

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

93

K. Omasa et al.

light image of the same region in B and C captured with acolor CCD camera. A stoma is at the center of the image, andthe epidermal cells have intricate shapes. Epidermal cellchloroplasts emitted more intense ﬂuorescence than themesophyll or guard cell chloroplasts (Fig. 3B, C). The guard
cell chloroplasts were smaller and denser than the mesophyll
chloroplasts. The inclination of the leaf to the right is apparent in the proﬁle image (Fig. 3B, C).
After the reconstructed 3D chlorophyll ﬂuorescence intensity images were processed by afﬁne transformation to adjust
the position gap between the images and to extract only
the chloroplasts images, the chlorophyll ﬂuorescence yields
and parameters such as ΦPSII and NPQ were calculated by
Equations (1)–(5) (see Materials and Methods). Fig. 4 shows
close-up 3D images of ΦF, ΦFm′ and ΦPSII reconstructed from aseries of chlorophyll ﬂuorescence intensity images captured
under 198 µmol m–2 s–1 PPF actinic light and 4,820 µmol m–2 s–1
PPF saturation pulse; the images were taken at the same site
shown in Fig. 3. Proﬁle images were reconstructed for theareas surrounded by the red and the sky-blue dotted lines(Fig. 4A) and are shown at the bottom and the right side,
respectively, of the abaxial surface image. Epidermal cellchloroplasts had larger ΦF and ΦFm′ values than guard celland mesophyll chloroplasts. The ΦPSII values of the epidermal cell chloroplasts and the guard cell chloroplasts werelower than those of the subjacent mesophyll chloroplasts.
3D-ΦF, 3D-ΦFm′ and 3D-ΦPSII images were reconstructed
clearly, because there was very little movement of all thechloroplasts during the capture of iF and iFm′ images, but
accurate 3D-NPQ images were not reconstructed, because
there was movement of some of the chloroplasts—especiallythe mesophyll chloroplasts—during the gap of several tensof minutes between the captures of iFm and iFm′.

FF
0.008

30 mm

30 mm

30 mm

0

A

0

Relationships between actinic light intensity andchlorophyll ﬂuorescence parameters of chloroplasts
in mesophyll cells, guard cells and epidermal cells
We examined the relationships between actinic light intensity and the chlorophyll ﬂuorescence parameters ΦPSII and
NPQ for the Boston fern and broad bean chloroplasts on theabaxial leaf side. As the actinic light intensity increased, ΦPSII
decreased and NPQ increased in all mesophyll, guard celland epidermal chloroplasts of Boston fern and broad beanleaf specimens. The ΦPSII values were lower for Boston fern
chloroplasts than for broad bean chloroplasts, and vice versa
for the NPQ values (Fig. 5). The maximum photochemicalyield of PSII, ΦFv/ΦFm, for attached mature leaves on theabaxial side, measured with a pulse amplitude modulation(PAM) 101 ﬂuorimeter, was approximately 0.55 for Boston
fern plants and approximately 0.65 for broad bean plants.
These ΦFv/ΦFm values were similar to the mean ΦPSII values
for mesophyll chloroplasts at 12 µmol m–2 s–1 actinic light.
There was no bleaching of leaf specimens during theexperiments.
For Boston fern leaf specimens, the mean ΦPSII values ofmesophyll, epidermal and guard cell chloroplasts decreased
from 0.52 to 0.16, from 0.42 to 0.11 and from 0.34 to 0.09,
respectively, with increasing actinic light intensity (Fig. 5A).
The ΦPSII values of the mesophyll chloroplasts were larger
than those of the epidermal and guard cell chloroplasts at all
actinic light intensities. The ΦPSII values of epidermal chloroplasts were larger than those of the guard cell chloroplasts at
all actinic light intensities. The mean NPQ values of mesophyll, epidermal and guard cell chloroplasts increased from
0.20 to 1.81, from 0.34 to 1.56 and from 0.36 to 1.94, respectively, with increasing actinic light intensity (Fig. 5B). The
NPQ values of the guard cell chloroplasts were larger than

FFm'

0.01

0

B

FPSII

0.2

C

Fig. 4 Close-up 3D images of ΦF, ΦFm′ and ΦPSII reconstructed from a series of chlorophyll ﬂuorescence intensity images captured at the samesite shown in Fig. 3 under 198 µmol m–2 s–1 PPF actinic light and a saturation light pulse of 4,820 µmol m–2 s–1 PPF. (A) ΦF; (B) ΦFm′; (C) ΦPSII.
Proﬁle images were reconstructed in the areas surrounded by the red and the blue dotted lines (A) and are shown at the bottom and the rightside, respectively, of the abaxial surface image.

94

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

3D CLSM of chlorophyll ﬂuorescence parameters

A 0.8

B 2.0
1.6

1.2
NPQ

FPSII

0.6

0.4

0.8
0.2
0.4
0.0

0

100

200

300

0.0

0

100

PPF (mmol m−2s−1)

200

300

PPF (mmol m−2s−1)

Fig. 5 Relationships between actinic light intensity (PPF) and the chlorophyll ﬂuorescence parameters (A) ΦPSII and (B) NPQ. Open squares,
triangles and circles represent the values for the subjacent mesophyll chloroplasts, epidermal chloroplasts and guard cell chloroplasts on theabaxial side, respectively, of Boston fern leaf specimens. Filled squares and circles represent the values for the subjacent mesophyll chloroplasts
and guard cell chloroplasts on the abaxial side, respectively, of broad bean leaf specimens. The ΦPSII and NPQ values were the averages for
16 chloroplasts, and the vertical bars represent the standard errors (n = 16), except in the case of Boston fern epidermal cells irradiated with
12 µmol m–2 actinic light, for which the average values were calculated for eight chloroplasts (n = 8).

100.0

FPSII × PPF (mmol m−2s−1)

those of the mesophyll and epidermal chloroplasts at all
actinic light intensities. The NPQ values of the mesophyll
chloroplasts were larger than those of the epidermal chloroplasts. The exception was at 12 µmol m–2 s–1 actinic light; at
this PPF, the mesophyll value was slightly lower than that of
the epidermis.
For broad bean leaf specimens, the mean ΦPSII values ofmesophyll chloroplasts and guard cell chloroplasts decreased
from 0.63 to 0.36 and from 0.73 to 0.36, respectively, withincreasing actinic light intensity (Fig. 5A). The ΦPSII values ofthe mesophyll chloroplasts were lower than those of theguard cell chloroplasts at all actinic light intensities. The
mean NPQ values of mesophyll chloroplasts and guard cellchloroplasts increased from 0.16 to 1.40 and from 0.15
to 1.27, respectively, with increasing actinic light intensity(Fig. 5B). The NPQ values of the mesophyll chloroplasts
were larger than those of the guard cell chloroplasts at all
actinic light intensities. Epidermal chloroplasts were notfound in the broad bean leaf specimens. The ΦPSII and NPQ
values of broad bean mesophyll and guard cell chloroplasts
showed the opposite trend to the equivalent chloroplasts inBoston fern specimens.
The value obtained by multiplying the PPF value of actinic
light by the ΦPSII value is an indicator of photosynthetic electron transport rate (ETR) and CO2 ﬁxation rate. We therefore examined the relationship between actinic light intensity(PPF) and ΦPSII × PPF values (Fig. 6). For Boston fern leaf
specimens, the ΦPSII×PPF values of mesophyll, epidermal
and guard cell chloroplasts increased from 6.3 to38 µmol m–2 s–1, from 5.2 to 27 µmol m–2 s–1 and from 4.2 to

80.0

60.0

40.0

20.0

0.0
0

100

200

300

PPF (mmol m−2s−1)

Fig. 6 Relationships between actinic light intensity (PPF) andΦ PSII × PPF values obtained from data in Fig. 5 . The symbols
correspond to those shown in Fig. 5. The vertical bars represent thestandard errors.

21 µmol m–2 s–1, respectively, with increasing actinic lightintensity. For broad bean leaf specimens, the ΦPSII × PPF
values of mesophyll and guard cell chloroplasts increased
from 7.8 to 88 µmol m–2 s–1 and from 8.9 to 88 µmol m–2 s–1,
respectively. These results indicate that the increases in ETR
and CO2 ﬁxation rates of Boston fern chloroplasts are lower
than those of broad bean chloroplasts. In addition, the initial
increase in these rates was larger in the order mesophyll, epidermal and guard cell chloroplasts in Boston fern specimens

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

95

K. Omasa et al.

and in the order guard cell and mesophyll chloroplasts inbroad bean specimens.
We also examined the relationships between the ΦPSII and
NPQ values of mesophyll chloroplasts and the ΦPSII and NPQ
values of epidermal and guard cell chloroplasts (Figs. 7, 8).
We found rough correlations, related to the equation y = x,
between the ΦPSII and NPQ values of mesophyll chloroplasts
and the values of epidermal and guard cell chloroplasts.
Changes in ΦPSII values from mesophyll chloroplasts were−0.03 to −0.09 (−15 to −30%) for Boston fern epidermal
chloroplasts, −0.07 to −0.18 (−34 to −51%) for Boston fern
guard cell chloroplasts and 0.00 to 0.10 (0 to 19%) for broadbean guard cell chloroplasts. In comparison, the changes in
NPQ values from mesophyll chloroplasts were −0.25 to 0.14
(67 to −18%) for Boston fern epidermal chloroplasts, 0.07 to
0.21 (7 to 77%) for Boston fern guard cell chloroplasts and−0.01 to −0.13 (−5 to −24%) for broad bean guard cellchloroplasts.

Discussion
Performance of the new CLSM system forapplication of the saturation pulse method
The saturation pulse method has been proposed as a powerful tool for assessing photosynthetic parameters (Genty et al.
1989, Bilger and Björkman 1990, Schreiber 2004). Application of the saturation pulse method using the CLSM system
requires measurement of the laser beam intensity on thespecimen. However, the diameter of the laser beam as viewed
through the objective lens focused on the specimen is muchsmaller than the measuring face of the PPF sensor. We
addressed this problem by developing a method for converting the laser beam focused on the measuring plane via an
objective lens to PPF by using a PPF sensor and a metal
A 0.6

pinhole disk 50 µm in diameter (see Materials and Methods).
In these experiments, laser power (mW) was converted to
PPF (µmol m–2 s–1) by the equation PPF = 11,311 × laser
power – 833.4. The equation revealed that the PPF of the
CLSM at maximum laser power (23.0 mW) for chlorophyll
ﬂuorescence measurements was extremely strong at approximately 260,000 µmol m–2 s–1.
The light intensity changes were controlled with twoelectromagnetic shutters installed in the system. The intensity was maintained at the dark or actinic light value and had
the capacity to change rapidly from dark or actinic lightintensity to saturation light pulse intensity and then backagain. The time taken to maximize the light intensity of thesystem was approximately 0.12–0.15 s (see Fig. 2), andthe intensity was minimized within 0.27 s. These are fasterthan the equivalent times taken by commercially available
PAM chlorophyll ﬂuorometers (e.g. WALZ, MINI-PAM andPAM-101).
In previous imaging studies of laser-induced chlorophyll
ﬂuorescence transients (Omasa 1998, Omasa and Takayama
2002), the laser beam irradiations pulsed with a scan frequency of 30 s–1; this was because PPF averages of
50–200 µmol m–2 s–1 caused transients similar to those of acontinuous xenon lamp. In comparison, the laser scan frequency of our rotating pinhole- (Nikpow disk) type CLSM
system was from 300 to 360 s–1, which is 10–12 times thefrequency obtained in previous studies. For this reason, we
concluded that the pulse of the laser beam had no effect on
the photochemical or non-photochemical activity measured
by the chlorophyll ﬂuorescence. The ﬂuctuations in laser
beam intensity measured every 30 ms by the EM-CCD
camera were very small (actinic light, ±1.5% and saturation
light pulse, ±0.45%, see Supplementary Fig. A). Consequently, the pulsed laser beam irradiated with the rotating
B

1.0

1:1

1:1

Guard cell FPSII

Epidermal FPSII

0.8
0.4

0.2

0.6
0.4
0.2

0.0
0.0

0.2
0.4
Mesophyll FPSII

0.6

0.0
0.0

0.2

0.4
0.6
Mesophyll FPSII

0.8

1.0

Fig. 7 Relationships between ΦPSII values of mesophyll chloroplasts and those of epidermal cell and guard cell chloroplasts, obtained from data
in Fig. 5A. (A) Relationship between ΦPSII values of mesophyll and epidermal chloroplasts. (B) Relationship between ΦPSII values of mesophyll
and guard cell chloroplasts. The symbols correspond to those shown in Fig. 5. The vertical and horizontal bars represent the standard errors.

96

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

3D CLSM of chlorophyll ﬂuorescence parameters

A 2.0

B 2.0
1: 1

1.2
0.8
0.4
0.0
0.0

1: 1

1.6
Guard cell NPQ

Epidermal NPQ

1.6

1.2
0.8
0.4

0.4

0.8
1.2
Mesophyll NPQ

1.6

2.0

0.0
0.0

0.4

0.8
1.2
Mesophyll NPQ

1.6

2.0

Fig. 8 Relationships between NPQ values of mesophyll chloroplasts and those of epidermal cell and guard cell chloroplasts, obtained from data
in Fig. 5B. (A) Relationship between NPQ values of mesophyll and epidermal chloroplasts. (B) Relationship between NPQ values of mesophyll
and guard cell chloroplasts. The symbols correspond to those shown in Fig. 5. The vertical and horizontal bars represent the standard errors.

pinhole Nipkow disk could be used to supply actinic lightand saturation pulses to the plants.
The saturation pulse intensity required to saturate photosynthetic electron transport differed with the intensity ofactinic light. A saturation pulse of >800 µmol m–2 s–1 in thedark and 1,300 µmol m–2 s–1 under 300 µmol m–2 s–1 actinic
light was sufﬁcient to saturate the photosynthetic electrontransport in mesophyll and guard cell chloroplasts in theabaxial leaf surfaces of Boston fern specimens (Fig. 1). This isbecause adequately photoactivated PSII requires stronger
light; the excess light energy is dissipated as heat (Bilger and
Björkman 1990, Noctor and Horton 1990). Furthermore, as
demonstrated in Fig. 1, the consistent trends obtained by
the modiﬁed CLSM system and the conventional system forthe relationships between chlorophyll ﬂuorescence yieldand PPF values and in the minimum intensities required tosaturate the photosynthetic electron transport validated
our estimations of laser beam intensity.
Imaging of ubiquitous chloroplasts in leaf tissues underhigh-resolution objectives requires the reconstruction of clear
3D images from the many images captured at different focal
planes. Our modiﬁed CLSM system has made it possible toreconstruct the 3D images, even when the objective is ×40
and the leaf tissue was inclined (Fig. 3). The precise reconstruction of 3D images requires at least 64 focal planes, especially when 3D deconvolution is applied. To calculate ΦPSII
and NPQ it is necessary to capture iFm and iFm′ images while
the photosynthetic electron transport is saturated; we therefore examined the saturation duration of the photosynthetic
electron transport of PSII. Chlorophyll ﬂuorescence intensities peak as electron acceptors such as QA (the primary quinone acceptor after PSII), QB (the secondary quinone acceptor
after PSII) and plastoquinone are reduced; thereafter, quenchingoccurs under continual light (Lazár 1999, Strasser et al, 2004).
The time taken to reach photochemical saturation was longer
in dark-adapted leaves than that in light-adapted leaves. This
result implies that it took longer to reduce fully QA that had
been adequately oxidized in the dark than QA that had
already been reduced by actinic light (Fig. 2). Saturation
duration was longer in the light than in the dark. Heat dissipation may have been increased by acclimatization to light;
therefore, the saturation light did not cause as rapid a changein ∆pH through the thylakoid membranes as would haveoccurred in the dark (Gilmore et al. 1998, Kramer et al. 1999,
Krause and Jahns 2004). The ﬂuorescence intensity peaked
within 0.2 s after initiation of saturation light pulse irradiation
and remained at a maximum level for at least 2.2 s (Fig. 2).
The piezo z-scan unit installed on the objective made it possible to capture the image from each different focal plane at
30 ms. It was possible to capture 64 images of iFm or iFm′ for3D reconstruction at different focal planes during the 1.92 ssaturation period. The saturation light pulse did not cause
photoinhibition and depress the photochemical yield of PSII,
even at the highest light intensity of 6,180 µmol m–2 s–1 for
3.5 s (see Supplementary Fig. B). The focal planes for reconstructing the 3D-iF image were captured under actinic lightat an exposure time of 700–1,000 ms, or 3000 ms with a PPF
of 12 µmol m–2 s–1, without changing the EM-CCD camera
digital gain values from those used for capturing the iFm andiF ′ images. The 64 focal planes were captured in 44.8–64 s,m
or 192 s with a PPF of 12 µmol m–2 s–1. The relatively long exposure time was possible because of the steady-state ﬂuorescence intensity conditions that occurred under the actinic
light.

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

97

K. Omasa et al.

Photosynthetic properties of chloroplasts inmesophyll cells, guard cells and epidermal cells
Chlorophyll ﬂuorescence intensity images of leaf tissue crosssection surfaces reveal the light distribution within the leaf
(Takahashi et al. 1994, Koizumi et al. 1998, Han et al. 1999,
Osmond et al. 1999, Pfündel and Neubohn 1999, Vogelmann
and Evans 2002, Evans and Vogelmann 2003). In comparison,
the new CLSM system makes it possible to capture proﬁleimages at a much higher resolution without the need for leaf
cross-sections, although the depth is limited to within
100 µm. The abaxial proﬁle images captured from Boston
fern leaf specimens (Fig. 3) suggest that there are brightchloroplasts near the leaf surface—especially epidermal chloroplasts, which were exposed to stronger light than were thesubjacent mesophyll chloroplasts. The high density of chloroplasts near the cell boundaries implies that large numbersof chloroplasts are required near intercellular spaces to allow
efﬁcient CO2 ﬁxation (Evans and von Caemmerer 1996).
With increasing actinic light intensity, the mean values of
ΦPSII decreased and the mean values of NPQ increased in thesubjacent mesophyll, guard cell and epidermal chloroplasts(Fig. 5). This trend was observed because the photosynthetic
efﬁciency decreases as a result of the increase in heat dissipation that occurs under strong light conditions (Goh et al.
1999, Lawson et al. 2002, Lawson et al. 2003). As anotherreason for this trend, there might be a deﬁciency in the
supply of CO2 for photosynthesis by chloroplasts in the cellswith increasing actinic light intensity (Terashima et al. 2001,
Lowson et al. 2002). In addition, the abaxial leaf surface (withstomata) touching the cover glass was in water and the
adaxial leaf surface (without stomata) was in air, thus
decreasing the CO2 supply to the chloroplasts. The meanvalues of ΦPSII and NPQ varied between species and cell type(Fig. 5). At all actinic light intensities the ΦPSII values werelower for Boston fern chloroplasts than for broad bean chloroplasts, and vice versa for NPQ values. The ΦPSII and NPQ
values of Boston fern chloroplasts changed largely in response
to a light intensity increase at low light intensity, whereas
the values for broad bean chloroplasts did not. The ΦPSII
values of Boston fern chloroplasts decreased in the ordersubjacent mesophyll, epidermal and guard cell chloroplasts,
and the NPQ values decreased in the order guard cell, subjacent mesophyll and epidermal chloroplasts, except at
12 µmol m–2 s–1 actinic light, when the NPQ value for mesophyll chloroplasts was just lower than that for epidermal
chloroplasts. The ΦPSII and NPQ values of broad been mesophyll and guard cell chloroplasts showed a trend opposite tothose of Boston fern chloroplasts. The ΦPSII × actinic light
PPF value, as an indicator of the photosynthetic ETR and
CO2 ﬁxation rate, increased with increasing actinic lightintensity (Fig. 6). However, the rates for Boston fern chloroplasts did not exceed half the rates observed for broad bean

98

chloroplasts, and the increased rates were a little above
50 µmol m–2 s–1 actinic light.
The ΦPSII values of guard cell chloroplasts have been previously reported to be indistinguishable from, or only slightly
lower (minimum of 79%) than, those of mesophyll chloroplasts (Baker et al. 2001) according to the variety of plant(Lawson et al. 2002, Lawson et al. 2003). In general, our results
support this tendency (see Fig. 7). However, the light intensity was attenuated in the leaf tissue; therefore, the overlying
chloroplasts were irradiated by stronger light than theunderlying chloroplasts. Consequently, if the chloroplasts
are of the same quality, the ΦPSII values should be lower and
NPQ values higher in the overlying chloroplasts than in theunderlying chloroplasts. This trend was observed for ΦPSII
proﬁles (Fig. 3).
Our calculated ΦPSII values for chloroplasts of Boston fern
leaves with lower ΦFv/ΦFm (=0.55) value were lower thanthose generally known in higher plants (Lawson et al. 2002,
Lawson et al. 2003). This might have been the result of inefﬁciency of photosynthetic electron transport in the ferns or
a severe deﬁciency in the supply of CO2 for photosynthesis
by chloroplasts in the cells (Terashima et al. 2001, Lawson
et al. 2002), because there were no stomata in the adaxial leaf
surface exposed to air, and the abaxial leaf surface (with stomata) touched the cover glass wetted with water. The lower
ΦPSII × actinic light PPF values for Boston fern chloroplastsalso suggest depression of the photosynthetic ETR and CO2
ﬁxation rate, although this indicator is inﬂuenced by the
proportion of incident photons absorbed by PSII and othermetabolic processes such as photorespiration, O2 (throughthe Mehler reaction) and nitrate assimilation (Baker andOxborough 2004). The ΦPSII values for Boston fern guard cellchloroplasts were lower than the equivalent values observed
for mesophyll chloroplasts: the differences were –0.07 to–0.18 (–34 to –51%)—larger than those reported for Polypodium vulgare and Tradescantia albifora by Lawson et al.
(2003). Boston ferns have a greater number of small chloroplasts in their guard cells than do other higher plants (Figs.
3, 4); this may increase light absorption capacity and reduce
PSII operating efﬁciency (Lawson et al. 2003). These ﬁndings
may explain the higher NPQ values observed for guard cellchloroplasts than for mesophyll chloroplasts in the Boston
fern (Figs. 5B, 8). Furthermore, the PSII operating efﬁciency
was lower for the guard cell chloroplasts than for the adjacent mesophyll chloroplasts (Baker et al. 2001), and also forthe mesophyll chloroplasts far away from the guard cells(Fig. 4).
The ΦPSII values were slightly higher (the maximum is19%) for guard cell chloroplasts of broad bean leaves with
the normal ΦFv/ΦFm value of 0.65 than for mesophyll chloroplasts, especially at low actinic light intensity. These results
are consistent with those of previous reports, which foundthat the photosynthetic O2 evolution rate is higher in broad

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

3D CLSM of chlorophyll ﬂuorescence parametersbean guard cells than in mesophyll cells (Shimazaki et al.
1982, Shimazaki 1989). In comparison, these ﬁndings do notagree with those reported by Lawson et al. (2003). It is likelythat the energy for opening the stomata is provided mainlyby the mitochondria and partly by the chloroplasts, although
the energy source may vary with the prevailing environment(Shimazaki et al. 1983, Vani and Raghavendra 1994, Shimazaki
et al. 2007) in higher plants, and the activity of ribulose bisphosphate carboxylase is lower in guard cell chloroplasts
than in mesophyll chloroplasts (Kopka et al. 1997, Vavasseur
and Raghavendra 2005). The high ΦPSII values of broad bean
guard cell chloroplasts may be related to the supply of ATP
and NADH required for the opening of stomata.
Inhibition of photosynthetic electron transport from PSII
to the cytochrome b6/f complex (e.g. by the action of DCMU)
causes an increase in chlorophyll ﬂuorescence intensity(Govindjee 1995). In contrast, ΦPSII and NPQ decrease owing
to the disappearance of non-photochemical quenching(Govindjee and Seufferheld, 2002) caused by gradients of pH
through thylakoid membranes (Gilmore et al. 1998, Kramer
et al. 1999, Krause and Jahns 2004). The chlorophyll ﬂuorescence intensities were higher for Boston fern epidermal
chloroplasts than for mesophyll chloroplasts, whereas the
ΦPSII and NPQ values were lower (Figs. 7, 8). The differencesin ΦPSII of epidermal chloroplasts compared with that ofmesophyll chloroplasts ranged from –0.03 to –0.09 (–15 to–30%); these differences were smaller than those observed
between guard cell chloroplasts and mesophyll chloroplasts.
These results suggest that photosynthetic electron transport
is inherently or extrinsically more inefﬁcient in epidermal
chloroplasts than in mesophyll chloroplasts. Both ΦPSII and
NPQ are depressed when chronic photoinhibition occurs(Osmond et al. 1994). The epidermal chloroplasts are closer
to the outside of the leaf than are the mesophyll chloroplasts; therefore, the former might have been subjected tostronger light than the latter. Consequently, chronic photoinhibition might have occurred in the epidermal chloroplasts in response to the strong light.

Materials and Methods
Plant material
Boston fern (N. exaltata L. ‘Bostoniensis’) plants were acclimatized in pots (10 cm diameter and 10 cm high) in an environmentally controlled growth chamber for 4 weeks before
the experiments. The pots were ﬁlled with artiﬁcial soil
(mixture of vermiculite and perlite, 2 : 1, v/v). The plants
were illuminated for 12 h each day with ﬂuorescent lights at
a PPF of 150 µmol m–2 s–1. Air temperature was 24.5°C during
the day and 23.0°C at night. Relative humidity was 70%
during the day and 90% at night. Broad bean (V. faba L.
‘Komasakae’) plants were grown for 6–7 weeks after being sownin pots ﬁlled with Kureha compost (KUREHA Co., Chuo,

Tokyo, Japan). The plants were illuminated for 13 h each daywith ﬂuorescent lights at a PPF of 250 µmol m–2 s–1. Air temperature was 26.0°C during the day and 22.0°C at night. Relative humidity was 60% during the day and 80% at night.
Boston fern and broad bean plants were watered daily with a
1 : 1,000 dilution of HYPONeX nutrient solution (HYPONeX
Japan Co., Setagaya, Tokyo, Japan). Fully expanded matureintact leaves were used for the experiments. The experiments
were performed in room environments of 25°C air temperature and 60% relative humidity. Whole-leaf specimens of
Boston fern plants, with a major leaf axis approximately
1–1.5 cm long, were used for CLSM investigations. For CLSM
investigations of broad bean plants, leaf specimens were cut
into approximately 1.5 cm squares immediately after being
cut from an individual plant and were then placed immediately on a wetted cover glass. The abaxial leaf surface touching the cover glass was wetted with water, and the top
surface was kept exposed to air.

Computer-aided system for 3D chlorophyll
ﬂuorescence imaging with CLSM
The computer-aided chlorophyll ﬂuorescence imaging systemwith modiﬁed high-speed rotating pinhole disk (Nipkow disk)
CLSM consisted of an inverted light microscope, a confocal
scanner unit, a 488 nm wavelength laser diode system, twotypes of cooled CCD camera, various controlling devices anda personal computer. The Nipkow disk confocal scanner unit
(modiﬁed version of model CSU-10, Yokogawa Electric, Co.,
Musashino, Tokyo, Japan) was installed between a highly
sensitive cooled EM-CCD camera (C9100-12, Hamamatsu
Photonics K.K., Hamamatsu, Shizuoka, Japan) and an inverted
light microscope (IX-71, Olympus Co., Ltd., Shinjuku, Tokyo,
Japan). A laser diode system (HPU50100-PFS-2, λ = 488 nm,
Yokogawa Electric Co., Musashino, Tokyo, Japan) was usedas the light source for CLSM. To remove reﬂected light andto measure chlorophyll ﬂuorescence, a long-pass ﬁlter(λ > 515 nm) was built into the confocal scanner unit (Omasa
and Konishi 2008).
A z-scan motor, which changed the position of the objective along the z-axis, was controlled by the motor controller(MAC 5000, Ludl Electronic Products Ltd., Hawthorone, NY,
U.S.A.) with 0.1 µm accuracy. For more precise and rapid
control of the objective position, a piezo z-scan unit (PIFOC,
Physik Instruments, Waldbronn, Germany) was used. The
working distance and the resolution of the piezo z-scan unit
were 100 µm and 10 nm, respectively. Use of the piezo z-scanunit made it possible to capture an image of each different
focal plane in 30 ms. The z-scan motor and the piezo z-scanunit were controlled with the computer. External andbuilt-in electromagnetic shutters added to the confocal
scanner unit were used for passing, blocking or dimming thelaser beam through neutral density (ND) ﬁlters. Both shutters

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

99

K. Omasa et al.

were controlled by our own software via a shutter controller
connected to the computer. The external shutter could beswitched to select between the aperture and an ND ﬁlter,
and the built-in shutter could be switched between apertureand no aperture. Timing of the control of the piezo z-scanunit, the shutters and the A/D conversion of the EM-CCD
camera was synchronized. A disk rotation controller was
newly added to adjust the rotation speed of the Nipkow disk
and thus eliminate Moiré fringes resulting from delicate gaps
in timing between the disk speed and A/D conversion.
Chlorophyll ﬂuorescence intensity images were captured
with the EM-CCD camera at resolutions of 512 horizontal × 512 vertical pixels × 14-bit gray level per frame, recorded
as 16-bit digital data on the main memory in the computer,and analyzed by commercial software (MetaMorph, Molecular Devices Inc., Toronto, Canada; ERDAS IMAGINE, Leica
Geosystems Geospatial Imaging LLC, Heerbrugg, Switzerland) and our own software. Transmitted light images of theleaves irradiated from above with a halogen lamp unit
(U-LH100L-3, TH4-100, Olympus Co., Ltd., Shinjuku, Tokyo,
Japan) were captured with a cooled color CCD camera(C5310, Hamamatsu Photonics K.K., Hamamatsu, Shizuoka,
Japan). The images were recorded on the computer via a
digital color video recorder (DSR-V10, Sony Co., Minato,
Tokyo, Japan) at 640 horizontal × 480 vertical pixels perframe with 24-bit resolution. An immersion objective (UApo,
×40, numerical aperture: 1.35, Olympus Co., Ltd., Shinjuku,
Tokyo, Japan) was used for all experiments.

Performance of computer-aided CLSM system
EM-CCD camera characteristics. The sensitivity of the EMCCD camera, from 1 to approximately 2,000 times, was controlled from the computer by setting digital gain values at
0–255. The ﬂuorescence intensities of a standard ﬂuorescentsheet were measured with the computer-aided CLSM system at 30 different laser intensities and digital gain values of50, 100 and 150 for the EM-CCD camera. The values revealed
a positive linear relationship between ﬂuorescence intensity
and the A/D conversion level at each gain value for A/D conversion levels in the range 500–14,500. For this reason, weused A/D conversion levels in the range 500–14,500.
Measurement of PPF on the measuring plane through an objective.
A metal pinhole disk (PA-50, Sigma Koki Co., Ltd., Sumida,
Tokyo, Japan) was used to measure the PPF of the laser
beam on the measuring plane (specimens or ﬂuorescentsheet) through an objective (×40). The disk was 9.5 mm inexternal diameter and within 0.2 mm thick, and had one
pinhole 50 µm in diameter. The measurement procedure
was as follows: ﬁrst, the beam exit of the glass ﬁber connected to the laser system was set 4 cm above the PPF sensor(LI-250, Licor Inc., Lincoln, NE, U.S.A.) to achieve adequate
uniformity in the laser beam intensity on the sensor face,100

the diameter of which was approximately 8 mm. The PPF
of the laser beam was measured through the cover glassof the specimen, which was set on top of the PPF sensor.
Secondly, the metal pinhole disk was installed between thesensor and the cover glass. The 50 µm pinhole was ﬁxed on
the center of the sensor face. The PPF of the laser beam wasmeasured through the cover glass and the pinhole. The scale
factor between PPFs was derived by these ﬁrst and second
procedures with and without the pinhole and was calculated
as 5,035 [standard error = 39 (n = 10)]. Finally, the PPF was
measured on the measuring plane of the CLSM systemthrough the objective, with the pinhole and the cover glasson the sensor unchanged. The true PPF on the measuring
plane of the CLSM system was calculated by multiplying themeasured PPF by the scale factor. The relationship between
laser power and calculated PPF was expressed as a linear
equation (y = 11,311x – 833.4, where x is the laser power(mW) and y is the calculated PPF (µmol m–2 s–1,) R2 = 1).
Control and measurement of laser beam intensity forthe saturation pulse method. The chlorophyll ﬂuorescenceparameters ΦPSII and NPQ were calculated by the saturation
pulse method, by producing a time sequence combinationof darkness, actinic light and saturation light pulse on themeasuring plane of the microscope, with external and builtin electromagnetic shutters added to the confocal scannerunit. The laser beam intensity required to achieve a saturation light pulse was ﬁrst determined by adjustment of thelaser power and the above-mentioned direct measurement
method using the metal pinhole disk. The actinic light intensity was adjusted by selection of the ND ﬁlter inserted into
the external electromagnetic shatter. The time course of thelaser beam intensity (PPF) on the measuring plane of the
microscope was estimated from changes in the ﬂuorescenceintensity of the standard ﬂuorescent sheet, as measured
with the EM-CCD camera connected to the CLSM system.
The A/D conversion values of ﬂuorescence intensity were
calibrated in accordance with the PPF values of the abovementioned direct measurements.
The electromagnetic shutters opened within 90 ms in thedark and within 60 ms under the actinic light conditions togenerate the saturation light pulse under constant laser
power. The shutters closed within 270 ms after the irradiation of specimens for approximately 2 s with a saturation
light pulse. The actinic light and the saturation light pulse
irradiated the specimens with uncertainties of −1.46 to+1.37% and −0.45 to +0.32% of the average intensities, respectively (see Supplementary Fig. A).
Each of the numerous pinholes on the rotating Nipkow
disk built into the confocal scanner unit had a diameter of
50 µm. The pinholes were gyroidally placed on the disk
250 µm apart. The rotational frequency was set at 25–30
rounds s–1 to eliminate interference patterns (Moiré fringes).

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

3D CLSM of chlorophyll ﬂuorescence parameters

The pattern of the pinhole arrangement was replaced every
30° revolution of the disk such that each point on the leaf
was irradiated at 300–360 times s–1. In this way, the lightintensity measured with the EM-CCD and PPF sensor (LI-250,
Licor Inc., Lincoln, NE, U.S.A.) was the average at the site irradiated with the laser beam at 300–360 times s–1.

Conﬁrmation of saturation light pulse intensity andduration of saturation of photosynthetic electrontransport
To determine the saturation light intensity, changes in ﬂuorescence yields were examined at different light pulse intensities for 2 s in darkness and light using the CLSM system anda conventional system. Microscopic chlorophyll ﬂuorescenceintensity images of the abaxial leaf surfaces were captured
with the CLSM system (×40 objective) at different intensities of laser light pulses. To provide the ﬂexibility necessary
to allow changes in the ratio of saturation light pulse intensity to actinic light intensity, the actinic light was applied
through a blue band-pass ﬁlter (470 nm < λ <495 nm, Olympus Co., Ltd., Shinjuku, Tokyo, Japan) from the halogen lamp
unit instead of the 488 nm diode laser. Change overs fromactinic light to laser pulses were performed by manuallyrotating a knob on the microscope. The leaf specimen was set
on a wetted cover glass (NEO, 24 × 32 mm, Matsunami Glass
Ind., Ltd., Kishiwada, Osaka, Japan) and used for the CLSM
system. After the leaf specimen had been maintained in thedark for at least 20 min, it was irradiated with 300 µmol m–2 s–1
actinic light over a 30 min period before the laser pulse irradiations were started. The laser light pulses were applied for2 s at 1 min intervals to avoid activation of the xanthophyll
cycle. The mean values of chlorophyll ﬂuorescence yield(chlorophyll ﬂuorescence intensity/light intensity) of guard
cell chloroplasts (>10) and mesophyll chloroplasts (>20) on
the abaxial leaf side in response to different light pulse intensities in the dark and under actinic light were computed. The
chlorophyll ﬂuorescence intensity images from the abaxial
surfaces of the same leaf specimens were then captured witha conventional chlorophyll ﬂuorescence imaging system(Omasa and Takayama 2003), which consisted of several
180 W metal halide light sources (LS-M180, Sumita Optical
Glass Inc., Urawa, Saitama, Japan), each equipped with ashort-pass ﬁlter (4-96, λ <600 nm, Corning Inc., Corning, NY,
U.S.A.), a heat-absorbing ﬁlter and optical ﬁbers; and a cooled
CCD camera (C5985-02, Hamamatsu Photonics K.K., Hamamatsu, Shizuoka, Japan) equipped with a long-pass ﬁlter(2-64, λ >640 nm, Corning Inc., Corning, NY, U.S.A.) Actinic
light at 300 µmol m–2 s–1 and light pulses of different intensities were supplied with the metal halide lamps. The averagechlorophyll ﬂuorescence yields of the abaxial leaf surface
measured with the conventional system were comparedwith those of guard cell chloroplasts and mesophyll chloroplasts measured with the CLSM system.
To investigate the duration of saturation of photosynthetic electron transport during saturation light irradiation,
the changes in intensity of chlorophyll ﬂuorescence for theabaxial leaf cells in response to the saturation light were
measured with the CLSM system under different actinic light
conditions. Before irradiation with the saturation light, leaf
specimens were adapted to the dark or to actinic light at 98
or 248 µmol m–2 s–1 for at least 30 min. Saturation light andactinic light were produced with the 488 nm diode laser andthe electromagnetic shutters with an ND ﬁlter providing
4.11% transmittance. This gave a saturation light intensity of
4,870 µmol m–2 s–1 for leaf specimens adapted to the dark,
and a saturation light intensity 24.33 times the actinic lightintensity for leaf specimens adapted to actinic light at 98 or
248 µmol m–2 s–1.

Reconstruction of 3D chlorophyll ﬂuorescenceintensity images and calculation of chlorophyll
ﬂuorescence parameters
To calculate the chlorophyll ﬂuorescence parameters ΦPSII
and NPQ by the saturation pulse method, we measuredchanges in the intensity of chlorophyll ﬂuorescence imagesof leaf specimens by the CLMS system during a time sequence
combination of dark, actinic light and saturation light pulse(see Supplementary Fig. A). To reconstruct the 3D-iFm
image, a series of chlorophyll ﬂuorescence intensity images
taken during saturation laser light pulses in the dark ( iFm)
was ﬁrst captured at 64 different focal planes for 1.92 s from
0.2 s after the start of irradiation. The ﬁrst focal plane was
located close to the abaxial leaf surface and the last focal
plane was at a depth of 80 µm in the direction of the abaxial
surface. Each image was captured at 30 ms intervals (exposure time of the EM-CCD) after a 1.25 µm movement of the
focal plane. Secondly, after the leaf specimen had been irradiated by the actinic laser light for 20 min, another series ofchlorophyll ﬂuorescence intensity images ( iF) was captured
at 64 different focal planes for 0.064t s at t ms intervals(t = exposure time) to reconstruct the 3D-iF image. Finally,
just after the capture of iF, a ﬁnal series of chlorophyll ﬂuorescence intensity images (iFm′) taken during the saturation
laser light pulse under actinic light was captured in the same
way as the iFm to reconstruct the 3D-iFm′. Dark current
images were captured at both exposure times in the dark.
The dark current images are represented by iD30 and iDt, andwere captured when exposure times were 30 ms and t ms,
respectively.
Fig. 9 shows the procedure used for 3D reconstruction
and the calculation of ΦPSII and NPQ. The original digital
images were ﬁrst processed by a 3 × 3 median ﬁlter to remove
the spike noise. The 3D reconstruction was performed

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

101

K. Omasa et al.

A series of original digital images

Median filtering (3 × 3)

3D deconvolution

3D reconstruction

FF imageFF’m image

FFmimage

1-bit binary image

Affine transformation

Extraction of chloroplasts

Calculation of NPQ

Calculation of FPSII image

Extraction of FPSII image

Fig. 9 Procedure used for 3D image reconstruction and calculation of ΦPSII and NPQ. A series of chlorophyll ﬂuorescence intensity images, iFm,
iFm′ and iF, were used for the original digital images.

with MetaMorph software to select the brightest focal
plane (z coordinate) at each x–y coordinate from the 64
median-processed images measured at different focal planes.
The reconstructed images were used to calculate the 3D-ΦPSII
image and NPQ values. These images were also deconvoluted to enhance contrast by using the nearest–neighbormethod of commercially available software (AutoDeblur/
AutoVisualize, AutoQuant Imaging, Inc., Troy, NY, U.S.A.)The ΦPSII images were calculated from the 3D-reconstructed, afﬁne-transformed, and undeconvoluted iF, iFm′,
iD , and iD images by the following equations, because30talthough the deconvoluted image was clearer, the A/D conversion level of the chlorophyll ﬂuorescence intensity was
inaccurate. The symbols Φ, I and a represent yield, irradiated

(3)
where
The ΦPSII images of chloroplasts alone were obtained bymultiplying the 3D-ΦPSII undeconvoluted image by a 1-bit binary
image of chloroplasts extracted from the deconvoluted
ﬂuorescence image with a clearer edge for each chloroplast,
and a false-color 3D-ΦPSII image was displayed.
The NPQ values were calculated from 3D-reconstructed,
afﬁne-transformed and undeconvoluted iFm, iFm′, iD30 and iDt
images in accordance with the following equations.
(4)

(1)
where
light intensity and the absorption coefﬁcient of the photosynthetic pigments.
(2)

102

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

(5)

3D CLSM of chlorophyll ﬂuorescence parameters

For ΦPSII and NPQ calculated by Equations (1) and(4), the effects of absorption by plant pigments, in the
direction of the depth for 3D imaging, are canceled byincluding the ﬂuorescence yields in the numerator and thedenominator.

Relationships between actinic light intensity andchlorophyll ﬂuorescence parameters of chloroplasts
in mesophyll cells, guard cells and epidermal cells
To investigate the relationships between actinic light intensity and the chlorophyll ﬂuorescence parameters of chloroplasts in mesophyll, guard and epidermal cells, 3D-iFm, 3D-iFm′,
and 3D-iF images were obtained at different actinic lightintensities. For Boston fern leaf specimens, a series of iFm
images used to reconstruct the 3D-iFm image was captured
at a saturation light pulse of 5,950 µmol m–2 s–1 after dark
adaptation for 1 h. Thereafter, the specimens were maintained
under 59 µmol m–2 s–1 actinic light for 30 min and the iF images
for the 3D-iF image were captured. The actinic light waschanged from 59 to 105, 152, 198 and 245 µmol m–2 s–1 ininterval steps of 15 min, and the iF images for the 3D-iF imageat each actinic light intensity were captured in turn. The
iF ′ images for the 3D-iF ′ images were captured during satmmuration light pulses of 1,430, 2,560, 3,690, 4,820 and
5,950 µmol m–2 s–1 just after the measurement of the iF
images, respectively. In addition, 3D-iF and 3D-iFm images
were captured in turn at 12 µmol m–2 s–1 actinic light andduring a saturation light pulse of 1,430 µmol m–2 s–1 after thesame periods of dark adaptation. For broad bean leaf specimens, 3D-iF and 3D-iFm images were measured only at 12,105
and 245 µmol m–2 s–1 actinic light. The procedures were similar to those of Boston fern leaf specimens. Sixteen chloroplasts within the guard cells, epidermal cells and subjacentmesophyll cells were selected from the 3D images, and themean values and standard errors for ΦPSII, NPQ andΦPSII × actinic light intensity (PPF) were calculated for eachcategory of chloroplasts, with the exception of Boston fern
epidermal cells irradiated with 12 µmol m–2 actinic light, forwhich eight chloroplasts were selected. The EM-CCD exposure time (t) for measuring iF images was 700–1,000 ms. For
cells irradiated with 12 µmol m–2 actinic light the t value was
3,000 ms. The ΦFv/ΦFm values of attached leaves were measured with a PAM 101 chlorophyll ﬂuorimeter (Waltz).

Supplementary data
Supplementary data are available at PCP online.

Acknowledgements
We thank Mr. Hideo Hirukawa for his technical support in
the development of the system. We also thank Dr. Kotaro

Takayama and Dr. Ryosuke Endo for their helpful
discussions.

References
Baker, N.R. and Oxborough, K. (2004) Chlorophyll ﬂuorescence asa probe of photosynthetic productivity. In Chlorophyll a Fluorescence.
A Signature of Photosynthesis. Edited by Papageorgiou,G.C. and
Govindjee, pp. 65–82. Springer, Dordrecht, The Netherlands.
Baker, N.R., Oxborough, K., Lawson, T.L. and Morison, J.I.L. (2001) High
resolution imaging of photosynthetic activities of tissues, cells andchloroplasts in leaves. J. Exp. Bot. 52: 615–621.
Bilger, W. and Björkman, O. (1990) Role of the xanthophyll cycle inphotoprotection elucidated by measurements of light-induced
absorbency changes, ﬂuorescence and photosynthesis in leaves ofHedera canariensis. Photosynth. Res. 25: 173–185.
Chaerle, L., Leinonen, I., Jones, H.G. and Van Der Straeten, D. (2007)
Monitoring and screening plant populations with combined thermaland chlorophyll ﬂuorescence imaging. J. Exp. Bot. 58: 773–784.
Daley, P.F., Raschke, K., Ball, J.T. and Berry, J.A. (1989) Topography ofphotosynthetic activity of leaves obtained from video images ofchlorophyll ﬂuorescence. Plant Physiol. 90: 1233–1238.
Endo, R. and Omasa, K. (2004) Chlorophyll ﬂuorescence imaging ofindividual algal cells: effects of herbicide on Spirogyra distenta at
different growth stages. Environ. Sci. Technol. 38: 4165–4168.
Endo, R. and Omasa, K. (2007) 3-D cell-level chlorophyll ﬂuorescenceimaging of ozone-injured sunﬂower leaves using a new passive lightmicroscope system. J. Exp. Bot. 58: 765–772.
Evans, J.R. and Vogelmann, T.C. (2003) Proﬁles of C-14 ﬁxation throughspinach leaves in relation to light absorption and photosynthetic
capacity. Plant Cell Environ. 26: 547–560.
Evans, J.R. and von Caemmerer, S. (1996) Carbon dioxide diffusion
inside leaves. Plant Physiol. 110: 339–346.
Genty, B., Briantais, J.M. and Baker, N.R. (1989) The relationshipbetween the quantum yield of photosynthetic electron-transportand quenching chlorophyll ﬂuorescence. Biochim. Biophys. Acta
990: 87–92.
Genty, B. and Meyer, S. (1995) Quantitative mapping of leaf
photosynthesis using chlorophyll ﬂuorescence imaging. Aust. J. Plant
Physiol. 22: 277–284.
Gilmore, A.M., Shinkarev, V.P., Hazlett, T.L. and Govindjee (1998)
Quantitative analysis of the effects of intrathylakoid pH andxanthophyll cycle pigments on chlorophyll a ﬂuorescence
lifetime distributions and intensity in thylakoids. Biochemistry 37:
13582–13593.
Goh, C.-H., Schreiber, U. and Hedrich, R. (1999) New approach ofmonitoring changes in chlorophyll a ﬂuorescence of single guard
cells and protoplasts in response to physiological stimuli. Plant Cell
Environ. 22: 1057–1070.
Govindjee (1995) Sixty-three years since Kautsky: chlorophyll a
ﬂuorescence. Aust. J. Plant Physiol. 22: 131–160.
Govindjee and Nedbal, L. (2000) Seeing is believing. Photosynthetica 38:
481–482.
Govindjee and Seufferheld, M.J. (2002) Non-photochemical quenchingof chlorophyll a ﬂuorescence: early history and characterization of
two xanthophyll-cycle mutants of Chlamydomonas reinhardtii.
Funct. Plant Biol. 29: 1141–1155.

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

103

K. Omasa et al.

Han, T., Vogelmann, T. and Nishio, J. (1999) Proﬁles of photosynthetic
oxygen-evolution within leaves of Spinacia oleracea. New Phytol.
143: 83–92.
Ichihara, A., Tanaami, T., Isozaki, K., Sugiyama, Y., Kosugi, Y., Mikuriya,
K., et al. (1996) High speed confocal ﬂuorescence microscopy using
a Nipkow scanner with microlenses for 3-D imaging of single
ﬂuorescent molecule in real-time. Bioimages 4: 57–62.
Koizumi, M., Takahashi, K., Mineuchi, T., Nakamura, K. and Kano, H.
(1998) Light gradients and the transverse distribution of chlorophyll
ﬂuorescence in mangrove and Camellia leaves. Ann. Bot. 81:
527–533.
Kopka, J., Provart, N.J. and Müller-Röber, B. (1997) Potato guard cells
respond to drying soil by a complex change in the expression ofgenes related to carbon metabolism and turgor regulation. Plant J.
11: 871–882.
Kramer, D.M., Sacksteder, C.A. and Cruz, J.A. (1999) How acidic is thelumen? Photosynth. Res. 60: 151–163.
Krause, G.H., Jahns, P. (2004) Non-photochemical energy dissipation
determined by chlorophyll ﬂuorescence quenching: characterizationand function. In Chlorophyll a Fluorescence. A Signature of Photosynthesis. Edited by Papageorgiou, G.C. and Govindjee, pp. 463–495.
Springer, Dordrecht, The Netherlands.
Küpper, H., Šetlίk, I., Trtίlek, M. and Nedbal, L. (2000) A microscope fortwo-dimensional measurements of in vivo chlorophyll ﬂuorescencekinetics using pulsed measuring radiation, continuous actinic
radiation, and saturating ﬂashes. Photosynthetica 38: 553–570.
Lawson, T., Oxborough, K., Morison, J.I.L. and Baker, N.R. (2002)
Responses of photosynthetic electron transport in stomatal guard
cells and mesophyll cells in intact leaves to light, CO2, and humidity.
Plant Physiol. 128: 52–62.
Lawson, T., Oxborough, K., Morison, J.I.L. and Baker, N.R. (2003) The
responses of guard and mesophyll cell photosynthesis to CO2, O2,
light, and water stress in a range of species are similar. J. Exp. Bot. 54:
1743–1752.
Lazár, D. (1999) Chlorophyll a ﬂuorescence induction. Biochim. Biophys.
Acta 1412: 1–28.
Leipner, J., Oxborough, K. and Baker, N.R. (2001) Primary sites of ozoneinduced perturbations of photosynthesis in leaves: identiﬁcationand characterization in Phaseolus vulgaris using high resolution
chlorophyll ﬂuorescence imaging. J. Exp. Bot. 52: 1689–1696.
Lichtenthaler, H.K. and Miehé, J.A. (1997) Fluorescence imaging as adiagnostic tool for plant stress. Trends Plant Sci. 2: 316–320.
Minsky, M. (1988) Memoir on inventing the confocal scanningmicroscope. Scanning 10: 128–138.
Nakano, A. (2002) Spinning-disk confocal microscopy—a cutting-edge
tool for imaging of membrane trafﬁc. Cell Struct. Funct. 27:
349–355.
Noctor, G. and Horton, P. (1990) Uncoupler titration of energydependent chlorophyll ﬂuorescence quenching and photosystem-II
photochemical yield in intact pea-chloroplasts. Biochim. Biophys.
Acta 1016: 228–234.
Omasa, K. (1998) Image instrumentation of chlorophyll a ﬂuorescence.
SPIE 3382: 91–99.
Omasa, K. (2000) 3-D color video microscopy of intact plants. In Image
Analysis: Methods and Applications, 2nd edn. Edited by Häder, D.P.
pp. 257–273 CRC Press, Boca Raton, FL.

104

Omasa, K., Hosoi, F. and Konishi, A. (2007) 3D lidar imaging fordetecting and understanding plant responses and canopy structure.
J. Exp. Bot. 58: 881–898.
Omasa, K., Konishi, A. (2008) Development of a 3D confocal laser
scanning microscope for applying the saturation pulse method tochlorophyll a ﬂuorescence. In Photosynthesis. Energy from the Sun.
14th International Congress on Photosynthesis. Edited by Allen, J.F.,
Gantt, E., Golbeck, J.H., and Osmond, B. pp. 657–660. Springer,
Berlin.
Omasa, K., Shimazaki, K, Aiga, I., Larcher, W. and Onoe, M. (1987) Image
analysis of chlorophyll ﬂuorescence transients for diagnosing
the photosynthetic system of attached leaves. Plant Physiol. 84:
748–752.
Omasa, K., Takayama, K. (2002) Image instrumentation of chlorophyll
a ﬂuorescence for diagnosing photosynthetic injury. In Air Pollution
and Plant Biotechnology. Edited by Omasa, K., Saji, H., Yousseﬁan, S.,
and Kondo, N. pp. 287–308, Springer, Tokyo.
Omasa, K. and Takayama, K. (2003) Simultaneous measurement of
stomatal conductance, non-photochemical quenching, andphotochemical yield of photosystem II in intact leaves bythermal and chlorophyll ﬂuorescence imaging. Plant Cell Physiol. 44:
1290–1300.
Osmond, C.B. and Grace, S.C. (1994) Perspectives on photoinhibitionand photorespiration in the ﬁeld: quintessential inefﬁciencies ofthe light and dark reactions of photosynthesis? J. Exp. Bot. 46:
1351–1362.
Osmond, B., Park, Y.-M. (2002) Field-portable imaging system formeasurement of chlorophyll ﬂuorescence quenching. In Air
Pollution and Plant Biotechnology. Edited by Omasa, K., Saji, H.,
Yousseﬁan, S., and Kondo, N. pp. 309–319. Springer, Tokyo.
Osmond, B., Schwarts, O. and Gunning, B. (1999) Photoinhibitory
printing on leaves, visualized by chlorophyll ﬂuorescence imaging
and confocal microscopy, is due to diminished ﬂuorescence from
grana. Aust. J. Plant Physiol. 26: 717–724.
Oxborough, K. (2004) Using chlorophyll a ﬂuorescence imaging tomonitor photosynthetic performance. In Chlorophyll a Fluorescence.
A Signature of Photosynthesis. Edited by Papageorgiou, G.C., andGovindjee. pp. 409–428. Springer, Dordrecht, The Netherlands.
Oxborough, K. and Baker, N.R. (1997a) An instrument capable ofimaging chlorophyll a ﬂuorescence from intact leaves at very low
irradiance and at cellular and subcellular levels of organization. Plant
Cell Environ. 20: 1473–1483.
Oxborough, K. and Baker, N.R. (1997b) Resolving chlorophyll a
ﬂuorescence images of photosynthetic efﬁciency into photochemical
and non-photochemical components—calculation of qP and Fv′/Fm′ measuring Fo′. Photosynth. Res. 54: 135–142.
Papageorgiou, G.C., Govindgee (eds) (2004) Chlorophyll a Fluorescence. A Signature of Photosynthesis. Springer, Dordrecht, The
Netherlands.
Pfündel, E. and Neubohn, B. (1999) Assessing photosystem I andphotosystem II distribution in leaves from C4 plants using confocal
laser scanning microscopy. Plant Cell Environ. 22: 1569–1577.
Rigaut, J.P., Carvajal-Gonzalez, S., Vassy, J. (1992) Confocal image
cytometry—quantitative analysis of three-dimensional images
obtained by confocal scanning microscopy. In Image Analysis
in Biology. Edited by Häder, D.P, pp. 109–133. CRC Press, Baca
Raton, FL.

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

3D CLSM of chlorophyll ﬂuorescence parameters

Rolfe, S.A. and Scholes, J.D. (1995) Quantitative imaging of chlorophyll
ﬂuorescence. New Phytol. 131: 69–79.
Rolfe, S.A. and Scholes, J.D. (2002) Extended depth-of-focus imaging ofchlorophyll ﬂuorescence from intact leaves. Photosynth. Res. 72:
107–115.
Schreiber, U. (2004) Pulse-amplitude-modulation (PAM) ﬂuorometry
and saturation pulse method: an overview. In Chlorophyll a
Fluorescence. A Signature of Photosynthesis. Edited by Papageorgiou,
G.C. and Govindgee. pp. 279–319. Springer, Dordrecht, The
Netherlands.
Schurr, U., Walter, A. and Rascher, U. (2006) Functional dynamics ofplant growth and photosynthesis—from steady-state to dynamics—
from homogeneity to heterogeneity. Plant Cell Environ. 29: 340–352.
Shimazaki, K. (1989) Ribulosebisphosphate carboxylase activity andphotosynthetic O2 evolution rate in Vicia guard-cell protoplasts.
Plant Physiol. 91: 459–463.
Shimazaki, K., Doi, M. Assmann, S.M. and Kinoshita, T. (2007) Light
regulation of stomatal movement. Annu. Rev. Plant Biol. 58: 219–247.
Shimazaki, K., Gotow, K. and Kondo, N. (1982) Photosynthesis
properties of guard cell protoplasts from Vicia faba L. Plant Cell
Physiol. 23: 871–879.
Shimazaki, K., Gotow, K., Sakaki, T. and Kondo, N. (1983) High
respiratory activity of guard cell protoplasts from Vicia faba L. Plant
Cell Physiol. 24: 1049–1056.
Strasser, R.J., Tsimilli-Michael, M., Srivastava, A. (2004) and Analysis of
the chlorophyll a ﬂuorescence transient. In Chlorophyll a
Fluorescence. A Signature of Photosynthesis. Edited by Papageorgiou,
G.C.,Govindjee. 321–362. Springer, Dordrecht, The Netherlands.
Takahashi, K., Mineuchi, K., Nakamura, T., Koizumi, M. and Kano, H.
(1994) A system for imaging transverse distribution of scattered

light and chlorophyll ﬂuorescence in intact rice leaves. Plant Cell
Environ. 17: 105–110.
Tanaami, T., Otsuki, S., Tomosada, N., Kosugi, Y., Shimizu, M. and Ishida
H. (2002) High-speed 1-frame/ms scanning confocal microscopewith a microlens and Nipkow disks. Appl. Opt. 41: 4704–4708.
Terashima, I., Miyazawa, S.I. and Hanba, Y. (2001) Why are sun leaves
thicker than shade leaves? Consideration based on analyses of CO2
diffusion in the leaf. J. Plant Res. 114: 93–105.
Tlałka, M. and Fricker, M. (1999) The role of calcium in blue-lightdependent chloroplast movement in Lemna trisulca L. Plant J. 20:
461–473.
Tlałka, M., Runquist, M. and Fricker, M. (1999) Light perception andthe role of the xanthophyll cycle in blue-light-dependent chloroplast
movements in Lemna trisulca L. Plant J. 20: 447–459.
Vani, T. and Raghavendra, A.S. (1994) High mitochondrial activity but
incomplete engagement of the cyanide-resistant alternative
pathway in guard cell protoplasts of pea. Plant Physiol. 105:
1263–1268.
Vavasseur, A. and Raghavendra, A.S. (2005) Guard cell metabolism andCO2 sensing. New Phytol. 165: 665–682.
Vogelmann, T.C. and Evans, J.R. (2002) Proﬁles of light absorption andchlorophyll within spinach leaves from chlorophyll ﬂuorescence.
Plant Cell Environ. 25: 1313–1323.
Wang, E., Babbey, C.M. and Dunn, K.W. (2005) Performance comparison
between the high-speed Yokogawa spinning disc confocal systemand single-point scanning confocal systems. J. Microsc. 218:
148–159.

(Received April 2, 2008; Accepted November 12, 2008)

Plant Cell Physiol. 50(1): 90–105 (2009) doi:10.1093/pcp/pcn174 © The Author 2008.

105