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TECHNICAL ADVANCE

Delayed fluorescence as a universal tool for the measurement

of circadian rhythms in higher plantsPeter D. Gould, Patrick Diaz, Claire Hogben, Jelena Kusakina, Radia Salem, James Hartwell and Anthony Hall *

School of Biological Sciences, University of Liverpool, Crown Street, Liverpool, UK

Received 11 November 2008; revised 7 January 2009; accepted 22 January 2009; published online 11 March 2009.*For correspondence (fax +44 151 795 4403; e-mail [email protected]).

SUMMARY

The plant circadian clock plays an important role in enhancing performance and increasing vegetative yield.

Much of our current understanding of the mechanism and function of the plant clock has come from the

development of Arabidopsis thaliana as a model circadian organism. Key to this rapid progress has been the

development of robust circadian markers, specifically circadian-regulated luciferase reporter genes. Studies of

the clock in crop species and non-model organisms are currently hindered by the absence of a simple high-

throughput universal assay for clock function, accuracy and robustness. Delayed fluorescence (DF) is a

fundamental process occurring in all photosynthetic organisms. It is luminescence-produced post-illumination

due to charge recombination in photosystem II (PSII) leading to excitation of P680 and the subsequent

emission of a photon. Here we reportthat theamount of DF oscillates with an approximately 24-h period andis

under the control of the circadian clock in a diverse selection of plants. Thus, DF provides a simple clock output

that may allow the clock to be assayed in vivo in any photosynthetic organism. Furthermore, our data provide

direct evidence that the nucleus-encoded, three-loop circadian oscillator underlies rhythms of PSII activity in

the chloroplast. This simple, high-throughput and non-transgenic assay could be integrated into crop breeding

programmes, the assay allows the selection of plants that have robust and accurate clocks, and possibly

enhanced performance and vegetative yield. This assay could also be used to characterize rapidly the role andfunction of any novel Arabidopsis circadian mutant.

Keywords: circadian, delayed fluorescence, Arabidopsis, chloroplast, photosystem II, luciferase.

INTRODUCTION

Thecircadian clock is an endogenous24-htimer that is found

in most eukaryotes and photosynthetic bacteria. The clock

plays an important role in the biology of an organism and

allows the synchronization of critical physiological, bio-

chemical and developmental processes with the local light/

dark (LD) cycle. In Arabidopsis thaliana (Arabidopsis), tran-

scriptomic studies have revealed the important role that the

clock plays in temporal organization and synchronization of

plant biology (Harmer et al., 2000; Edwards et al., 2006).

Important agronomic traits, such as water use efficiency and

photoperiodic control of flowering time, are regulated by the

clock. Furthermore, plants with dysfunctional clocks have

reduced water use efficiency, dry weight and photosynthetic

CO2 fixation, which supports the long-held theory that the

clock contributes to a plant’s ‘fitness’ (Dodd et al., 2005). The

next clear goal in plant circadian biology is to investigate

whether robust and accurate clock function is an important

‘fitness’ trait for crop species. One fundamental limitation to

achieving this goal is the lack of a simple and high-through-

put method for assaying clock function across species.

At present, much of our understanding of the plant

circadian clock has come from the use of Arabidopsis as a

model circadian organism. Critical to this use has been the

development of a robust high-throughput assay for mea-

suring rhythms in this plant that utilizes clock-controlled

promoter luciferase (LUC) reporter fusions (Millar et al.,

1992). While luciferase could offer a universal assay for clock

function, and has been used to measure rhythms in rice

ª 2009 The Authors 893Journal compilation ª 2009 Blackwell Publishing Ltd

The Plant Journal (2009) 58, 893–901 doi: 10.1111/j.1365-313X.2009.03819.x



(Sugiyama et al., 2001), tobacco and Arabidopsis (Millar

et al., 1992), it requires the insertion of promoter:LUC

fusions into the plant genome, a technique only suited to

species in which stable transformation is routine.

Alternative methods to the luciferase technique do exist,

such as the measurement of rhythmic patterns in growth ororgan movement. For instance, circadian rhythms in leaf

movement have been described in a numberof dicot species

including Arabidopsis, Phaseolus (Bu ¨ nning, 1935) and Bras-

sica oleracea (Salathia et al., 2007). However, in monocots,

there has been only one report of a growth/movement

rhythm; namely the extension of the coleoptile of Avena

sativa seedlings in constant darkness (DD) (Ball and Dyke,

1954). Another potential universal assay of the clock is an

infra-red gas analyser (IRGA) system for the measurement of

CO2 assimilation and stomatal conductance. However, due

to the complexity of these systems only low-throughput

versions exist. The highest capacity reported for a multi-

channel IRGA gas exchange system is six channels (Dodd

et al., 2004). There is a real need for the development of a

robust, high-throughput method that does not require

production of transgenic plants and can be used to assay

clock function in important crop species.

Delayed fluorescence (DF) or delayed light emission was

discovered in 1951 by Strehler and Arnold (Strehler and

Arnold, 1951), it is a well-studied and fundamental process

found in all photosynthetic organisms (reviewed in: Jursinic,

1986). It results from the post-illumination emission of light

from chlorophyll a, principally from photosystem II (PSII), as

a result of charge recombination between excited plasto-

quinone QA and P680 leading to the emission of a photon(Rutherford et al., 1984). Delayed fluorescence, therefore,

offers a simple method of probing PSII photochemistry.

Critically, with the advent of photomultiplier tubes and low

light imaging cameras, DF is a simple measurement to

make. We demonstrate here that it can be measured using a

charged coupled device (CCD) camera system that has been

developed for the in vivo monitoring of promoter:LUC

activity (Southern et al., 2006).

Here we show that thelevel of DF is under robustcircadian

control and provides a simple assay for measuring period,

robustness and accuracy of the circadian clock. Using well

characterized Arabidopsis circadian clock mutants, we dem-

onstrate that rhythms in DF are controlled by the same

molecular oscillator that drives rhythms in other circadian

outputs. We show that DF may provide a universal method

for measuring circadian rhythms in higher plants, demon-

strating robust rhythms in Capsella bursa-pastoris (C. bursa-

pastoris ), Lactuca sativa (lettuce), Hordeum vulgare (barley),

Zea mays (maize) and Kalanchoe fedtschenkoi (K. fed-

tschenkoi ). This method offers a simple, high-throughput

way of measuring circadian rhythms using existing technol-

ogy and does not require the insertion of a reporter

transgene.

RESULTS

Measuring DF in Arabidopsis

The measurement of DF requires immediate removal of

actinic light, followed by detection of the weak DF signalusing a highly sensitive detector. These requirements are all

similar to those for luciferase imaging (Southern et al.,

2006). We used our existing luciferase imaging system to

measure DF. The system was modified slightly, removing or

covering any auto-fluorescent materials in the imaging

chamber. The light source consisted of red and blue (RB)

light emitting diodes (LEDs) that did not auto-fluoresce

(Figure S1). The auto-fluorescent LED circuit boards were

sprayed black, and an electronic baffle circuit was included

to ensure rapid switching off of the LEDs. Using this system,

we were able to use a 1-min exposure to capture a 2D

luminescent image from a single wild-type (WT) Arabidopsis

seedling immediately after illumination. To investigate

whether the luminescence we were measuring was consis-

tent with previously described observations of DF, we first

measured decay kinetics of the luminescence. Light grown

Arabidopsis seedlings were placed in the imaging chamber

for 10 min under 35 lmol m)2 sec)1 of light. Immediately

after the lights were switched off, a time series of 10-sec

images with 100-msec delays between images was cap-

tured. Luminescence decayed rapidly and was undetectable

with our camera within 50 sec (Figure 1a). This rapid decay

was consistent with previous reports of DF decay kinetics

(Jursinic, 1986). To measure the emission spectra of the DF,

Figure 1. Both the spectral emission and kinetics of the DF response are

consistent with the emission of light from chlorophyll a.

(a) Decay kinetics of DF immediately after illumination. Each point of the

series represents the average integrated luminescence of 15–20 seedlings

during a 10-sec exposure.

(b) The wavelength of the emitted light immediately after lights off measured

in 9-day-old Arabidopsis seedlings. Each point represents the average

integrated luminescence of 15–20 seedlings during a 1-min exposure. The

series represents DF of the same seedlings measured from 400–720 nm in

20 nm steps.

894 Peter D. Gould et al.

ª 2009 The AuthorsJournal compilation ª 2009 Blackwell Publishing Ltd, The Plant Journal , (2009), 58, 893–901



a tuneable filter was attached to the front of the camera lens.

The spectral emission from the seedlings was measured

from 400 nm through to 720 nm in 20 nm steps. An emis-

sion peak was observed between 700 and 720 nm, consis-

tent with theDF beingproducedby chlorophyll a (Figure 1b).

The level of DF is under circadian control in WT Arabidopsis

A substantial body of published evidence supports the

hypothesis that the circadian clock controls photosynthesis

in plants. In Arabidopsis, microarray experiments have

revealed that many of the key genes that make up the light

harvesting complex and PSI and II are under circadian con-

trol at the level of the associated steady-state transcript

abundance (Harmer et al., 2000). There is also clear evidence

in multiple species that CO2 assimilation and light-induced

electron flow are clock controlled (Lonergan, 1981; Hennes-

sey and Field, 1991). In the marine dinoflagellate Gonyaulax

polyedra , rhythms in photosynthesis have been attributed to

PSII (Samuelsson et al., 1983). Chloroplast circadian period

has been linked to the circadian clock in the nucleus using a

chloroplast-targeted psbD:lucCP luciferase reporter strain of

the unicellular alga Chlamydomonas reinhardtii (Matsuo

et al., 2006). Furthermore, prompt chlorophyll fluorescence

imaging has been used to measure rhythms of photosyn-

thetic efficiency (/PSII) in leaves of the CAM plant Kalanchoe

daigremontiana (Rascher et al., 2001). Collectively, this evi-

dence led us to test whether: (i) the amount of light emitted

from a leaf during DF is subject to circadian control; (ii) DF

could be used as a non-invasive read-out of circadian

changes in PSII charge recombination; and (iii) DF could beused as a robust high-throughput method to assay clock

function.

To test for circadian rhythms in DF, Arabidopsis seedlings

from three different accessions (Col-0, Ws and C24) were

grown in groups of 15–20 seedlings in 12 h light/12 h dark

cycles at 22°C for 9 days on Murashige and Skoog (MS)

medium with 3% sucrose. The entrained seedlings were

then transferred at dawn to constant RB light (35 lmol m)2

sec)1) and temperature (22°C). This methodology was

consistent with that used for luciferase imaging, thus,

allowing comparison of luciferase and DF data. DF wasassayed every hour by switching the LED lights off and then

taking a 1-min exposure. This fully automated process was

repeated every hour for 96 h. The amount of luminescence

for each group of seedlings was corrected for background

and normalized as described in the methods (Figure 2). The

data clearly demonstrate that the clock drives robust

rhythms in the amount of DF in Arabidopsis ecotypes Ws,

Col-0 and C24, with a period of 23.3 h SE 0.1 n = 15, 24.5 h

SE 0.1 n = 7 and 24.1 h SE 0.4 n = 8 respectively. These

period estimates match closely with those for CHLORO-

PHYLL A/B BINDING PROTEIN 2:LUC (CAB2:LUC ) expres-

sion, a well characterized clock regulated gene (Table 1) and

leaf movement (Edwards et al., 2005) under similar exper-

imental conditions. However, the peak phase of DF for

Arabidopsis is approximately 2 h after dusk, whereas for

CAB2:LUC the peak phase of expression is approximately

4 h after dawn. The rhythms in DF also had higher relative

amplitude error (RAE) than those associated with rhythms in

CAB2:LUC expression and leaf movement rhythms. RAE is a

measure of rhythmrobustness varyingfrom 0 (a perfect fit to

the cosine wave) to 1 (not statistically significant). We have

also observed similar DF rhythms in single seedlings

(Figure S2a) and excised leaves of mature Arabidopsis

plants (Figure S2b). These observations are consistent with

clock regulation of the amount of DF. The similarity in theperiod of oscillation for DF with other circadian outputs in

Arabidopsis supports the hypothesis that DF rhythms are

driven by the same molecular oscillator. The rate of decay of

DF is unaffected by the clock; it is the absolute amount of DF

that is clock regulated (Figure S3). Using this assay, we can

Figure 2. The amount of DF is under circadian control.

Arabidopsis seedlings were grown on Murashige and Skoog basal salt mixture (MS) media containing 3% sucrose and entrained under 12 h light/12 h dark cycles

for 9 days at 22°C before transferring to constant RB light (35 lmol m)2 sec–1) for imaging.

(a) The plots represent normalized averages for DF of 7–15 groups of seedlings assayed in constant RB light every 1 h for 96 h. Ws n = 15 (black squares), C24 n = 8

(grey squares) and Col-0 n = 7 (empty squares). Error bars indicate SE.

(b) Period estimates for groups of seedlings plotted against their RAE. Ws n = 15 (black squares), C24 n = 8 (grey squares) and Col-0 n = 7 (empty squares).

Circadian rhythms in delayed fluorescence 895




measure rhythms in 400 single seedlings, or 256 groups

of seedlings, making the method high-throughput and

amenable to genetic screens.

The DF oscillator responds to the CCA1/LHY-TOC1/GI-

PRR7/9 nucleus-encoded clock

One intriguing question is whether photosynthetic rhythms

in the chloroplast are generated by the same mechanism

that drives rhythmic expression of transcription in thenucleus. DF provides a simple assay not only for the

circadian clock, but specifically for the circadian output from

PSII in the chloroplast. In Arabidopsis, our current model of

the central clock consists of a series of interlocking feedback

loops (Locke et al., 2006). One loop consists of TIMING OF

CAB EXPRESSION 1 (TOC1) and two closely related

Myb-transcription factors LATE ELONGATED HYPOCOTYL

(LHY ) and CIRCADIAN CLOCK ASSOCIATED 1 (CCA1)

(Alabadi et al., 2001). TOC1 forms a second loop with

GIGANTEA (GI ), and a third loop is formed between LHY/

CCA1 and the TOC1 paralogues PSEUDO-RESPONSE REG-

ULATOR 7 (PRR7 ) and PRR9 (Farre et al., 2005).

To investigate the question of whether circadian rhythms

of charge recombination in PSII are generated by the three-

loop oscillator and to demonstrate the utility of the assay for

the characterization of mutants we analysed the circadian

regulation of DF in Arabidopsis circadian clock mutants.

Previous analysis had identified that each one of these

mutants causes either period lengthening: prr7 , prr9 (Farre

et al., 2005), or shortening: cca1, lhy (Mizoguchi et al.,

2002), toc1 (Strayer et al., 2000), and gi (Park et al., 1999),

of all circadian outputs tested. Both mutant and WT plants

were grown under 12 h light/12 h dark cycles at 22°C for

9 days before transferring to constant RB light and 22°C.

The free running rhythms were plotted and periods mea-

sured for DF (Figures 3 and S4, and Table 1). The effect of

these mutations on the DF rhythms closely match those of

previously published CAB2:LUC data (Park et al., 1999;

Strayer et al., 2000; Mizoguchi et al., 2002; Farre et al.,2005). The close correlation between the effects of these

clock mutations on rhythms of DF and their effects on

CAB2:LUC expression supports the conclusion that both DF

and promoter activities of CCR2:LUC and CAB2:LUC are

driven by the same molecular oscillator. Thus, rhythmicity

in the chloroplast is generated by the same three-loop

oscillator. Furthermore, this experiment provides clear

evidence that even though the rhythms in DF are not as

robust as those for CAB2:LUC expression or leaf movement,

they can be successfully used to analyse subtle perturba-

tions of the circadian clock.

Rhythms in DF can be measured in a range of plant species

We investigated whether the rhythm of DF is unique to

Arabidopsis, or whether it could be observed in a diverse

range of plant species. We tested important monocotyle-

donous cereal crop species: barley, a C3 plant and maize a

C4 plant and a number of dicotyledonous species including

lettuce, C. bursa-pastoris and K. fedtschenkoi a model CAM

species (Figures 4 and S5). Importantly, rhythms could be

identified in all species; however some were more robust

than others. K. fedtschenkoi , maize and lettuce in particular

displayed robust and high-amplitude rhythms. For K. fed-

tschenkoi and maize time-lapse videos demonstrate robustoscillations in total DF (Figures S6 and S7). Furthermore, it

was clear that DF was not uniform across leaves with waves

of DF tracking over the leaf creating patchiness or hetero-

geneity consistent with that reported previously for prompt

fluorescence in K. daigremontiana (Rascher et al., 2001).

This assay offers a simple, high-throughput method for

assaying clock function in a range of important plant spe-

cies, including species in which no robust assay for clock

function currently exists.

Dual measurement of two circadian outputs

The luminescent properties of luciferase and DF are differ-

ent. Firstly, the two forms of bioluminescence produce light

of different wavelengths with firefly luciferase emitting light

at 560 nm and DF at 720 nm (Figure 1b). Secondly, upon

transfer to the dark the immediate amount of DF is large and

rapidly decays within 50 sec (Figure 1a), whilst, over a short

time scale (1 min), the luminescence from a clock pro-

moter:LUC reporter gene is low and fairly constant. Given

these different dynamic characteristics we investigated the

possibility that these two forms of bio-luminescence could

be simultaneously measured in the same plant.

Table 1 Period measurement for DF and CAB2:LUC rhythms from

Arabidopsis circadian clock mutants and their respective WT

ecotypes

Line

DF data LUC data

Period (h) SE (n ) Period (h) SE (n )

Ws 24.6 0.3 (16) 23.3 0.1 (15)

gi-11 24.2 0.5 (15) 22.8 0.2 (26)

lhy-21 23.5 0.2 (16) 21.6 0.1 (12)

cca1-11 23.6 0.2 (16) 21.2 0.1 (11)

C24 25.1 0.4 (9) 24.1 0.1 (8)

toc1-2 23.2 0.2 (16) 22.1 0.1 (16)

Col-0 24.4 0.1 (8) 24.3 0.1 (8)

prr7-3 25.5 0.2 (16) ND ND

prr9-1 25.4 0.16 (16) 24.6 0.2 (16)

Periods given are the variance-weighted means (period) of the

estimates for n groups, with variance-weighted standard errors of

the mean (SE). The luciferase data in this table are from CAB2 :LUC

rhythms produced in this lab. Differences in periods between WT and

all mutants are statistically significant (Student’s t -test P < 0.01). Thedata in this table are from a single representative experiment.





To achieve this dual imaging, seedlings transformed withCAB2:LUC were entrained under 12-h light/12-hdark cycles at

22°C for 9 days. The day before transferring to constant RB

light the plants were sprayedwith 5 mM luciferin. Plantswere

then transferred to constant RB light at dawn of thenext day.

To allow the capture of DF and luciferase data from a single

plant the imaging protocol had to be modified (see Experi-

mental procedures). This protocol allowed the capture of DF

and luciferase data every 2 h fora total of 96 h (Figure 5). The

phase, amplitude and periods of the DF (23.8 h, SE 0.2,

n = 16) and CAB2:LUC (24.2 h, SE 0.1, n = 16) rhythms were

similar to those measured using single assays. The differ-

ences in luminescent properties of DF and luciferase can

therefore be used to measurethese two rhythms in thesame

plant essentiallysimultaneously. Thisdual assay will provide

a useful tool to probe the mechanism by which the nuclear

oscillator drives rhythms in the chloroplast.

DISCUSSION

We have demonstrated circadian regulation of DF rhythms

for a number of crop and model plant species. Our charac-

terization of DF rhythms in a series of key circadian clock

mutants provides compelling evidence that the DF rhythms,

and hence chloroplastic rhythms, are driven by the nuclearencoded three-loop molecular oscillator. Taken together,

our results support the potential of DF as a universal, high-

throughput method of assaying central clock function in

higher photosynthetic organisms.

What is not clear from our results is the mechanism by

which DF is coupled to the molecular oscillator. One clear

difference between the rhythms in DF from different species

was the rhythm in K. fedtschenkoi , for which the period was

markedly shorter (Figure 4c). It is particularly noteworthy

that there was a good correlation between the period length

of the K. fedtschenkoi DF rhythm and the period of the CO2

fixation rhythm of equivalent leaves (Anderson and Wilkins,

1989). The close correlation suggests that the DF rhythm is a

reliable readout of the same oscillator that drives the CAM

circadian rhythm of CO2 fixation.

It has been suggested previously that the clock mecha-

nism driving rhythms in the chloroplast may be separate

from the well characterized nuclear clock (Roenneberg and

Hastings, 1988). Clear evidence in support of this comes

from the green macro-alga, Acetabularia. When the nucleus

is removed from these cells, rhythms still persist in the

chloroplast (Sweeney and Haxo, 1961). The DF we have

assayed is a measure of the photochemical state of PSII;

Figure 3. Arabidopsis circadian clock mutations

affect DF rhythms.

Arabidopsis seedlings were grown on MS media

containing 3% sucrose and entrained under 12-h

light/12-h dark cycles for 9 days at 22°C before

transferring to constant RB light (35 lmol m)2 -

sec)1) and assaying DF with 1-h time resolution

for 96 h.(a–c) The plots represent normalized averages

for DF of 8 to 16 groups of seedlings. (a) Ws

n = 16 (black squares), cca1-11 n = 16 (red

squares), lhy-21 n = 16 (yellow square) and gi-

11 n = 15 (green square); (b) Col-0 n = 8 (black

squares), prr7-3 n = 16 (red squares) and prr9-1

n = 16 (green squares); (c) C24 n = 9 (black

squares) and toc1-2 n = 16 (red squares). Error

bars indicate SE.

(d–f) Period estimates for groups of seedlings

plotted against their RAE. (d) Ws n = 16 (black

squares), cca1-11 n = 16 (red squares), lhy-21

n = 16 (yellow squares) and gi-11 n = 15 (green

squares). (e) Col-0 n = 8 (black squares), prr7-3

n = 16 (red squares) and prr9-1 n = 16 (green

squares). (f) C24 n = 9 (black squares) and toc1-2

n = 16 (red squares).





therefore, in measuring rhythms in DF, we were directly

assaying a clock-controlled output in the chloroplast. Here,

we have demonstrated that mutations in key clock compo-

nents affect rhythms in DF in an identical way to other

rhythmic outputs (Figure 3, Table 1). These data provide

strong evidence that rhythms in the chloroplast are driven

by the same oscillator driving other outputs. One intriguing

question arising from our work is: how does the nuclear

oscillator transduce its rhythmic output to the chloroplast?

One possible mechanism is via circadian regulated control

of intracellular calcium levels (Johnson et al., 1995). Calcium

has been implicated in both the assembly and function of

PSII (Brand and Becker, 1984). It is possible that calcium

rhythms could couple the rhythms in DF to the clock.

However, while cytosolic calcium levels oscillate in constant

light (LL), calcium does not oscillate in the chloroplast

(Johnson et al., 1995). It is, therefore, still unclear how

nuclear rhythms are transduced to the chloroplast. The DF

assay should provide an important tool for probing this

coupling mechanism.

As a circadian marker, DF has a number of distinct

advantages over current methods for assaying clock func-

tion. It does not require the insertion of a transgene and can

be used to assay clock function in plants that are difficult to

transform. Even for Arabidopsis, insertion of a transgene

either by crossing, or by transformation, can take

4–6 months. However, DF can be readily used to screen

existing mutant collections. It uses similar equipment to that

currently used to measure luciferase activity. It is a high-

throughput method and allows measurements of rhythms

from 400 single seedlings (Figure S2), or 256 groups of

seedlings (Figure 3) in a single experiment. Unlike leaf

movement rhythms, it allows accurate measurement of

phase as well as period.

Figure 4. DF rhythms can be measured in a

range of plant species.

Plants were entrained under 12-h light/12-h dark

cycles at 22°C before transferring to constant RB

light (35 lmol m)2 sec)1) and assaying DF with a

1-h time resolution for 96 h.

(a–e) The plots represent normalized averages

for DF of 8 to 42 groups of seedlings/leaves. (a)Lettuce n = 20; (b) maize n = 8; (c) K. fed-

tschenkoi n = 15; (d) barley n = 42 (j).

(e) C. bursa-pastoris n = 8. Error bars indicateSE.

(f–j) Period estimates plotted against their RAE.

(f) Lettuce n = 20; (g) maize n = 8; (h) K. fed-

tschenkoi n = 15; (i) barley n = 42; (j) C. bursa-

pastoris n = 8.





It has been reported previously that having a robust and

accurate clock increases photosynthesis and productivity in

Arabidopsis (Dodd et al., 2005). Measuring clock accuracy

and robustness in crop species, and correlating this with

yield and performance, will be essential when assessing

whether clock function is an important agricultural trait.

Here, we have demonstrated that DF can be used to measure

rhythms in a diverse range of species, thus providing a

powerful tool for investigating the correlation between clock

function and performance in crops.

EXPERIMENTAL PROCEDURES

Plant material

The T-DNA insertions cca1-11 and lhy-21 mutants were previously

described in (Hall et al., 2003). The gi-11 mutant was isolated from

T-DNA insertion lines and described in (Fowler et al., 1999). The

toc1-2 mutant was isolated in a clock mutant screen and was first

described in (Strayer et al., 2000). The T-DNA insertion mutants in

prr7-3 and prr9-1 were described in (Farre et al., 2005). The CA-

B2:LUC transgene, in the Ws background, was as described by Hall

(Hallet al.

, 2002).

Growth conditions

For Arabidopsis, lettuce and C. bursa-pastoris , seeds were surface

sterilized in 70% ethanol immediately followed by 50% bleach for

10 min. To remove traces of bleach they were rinsed with sterile

distilled water (SDW) and then re-suspended in 0.01% agar. The

seedlings were then sown on MS media, containing 3% sucrose

and 1.5% agar, in small clusters of 15–20 seeds. Seeds were kept

at 4°C in the dark for 2 days and then grown in 12-h light/12-h

dark cycles in a plant growth room at 22°C and 80 lmol m)2 sec)1

of light. Maize, barley and K. fedtschenkoi were grown on John

Innes No. 3 in identical growth conditions to the Arabidopsis

seedlings.

Measurement of DF

The imaging system for DF was identical to the luciferase system

described previously in (Southern et al., 2006). DF was detected

using an ORCA-11-BT 1024 16-bit low light charged coupled device

(CCD) camera cooled to )80°C (Hamamatsu Photonics; http://

www.hamamatsu.com). Attached to the camera was a high-

transmission lens (Xenon 0.95/25 mm, Schneider; http://www.schneiderkreuznach.com). The camera was inserted through a

modified port on the top of a Sanyo MIR-553 programmable cooled

incubator (Sanyo Gallenkamp; http://www.sanyo-biomedical.co.

uk), allowing precise temperature control. The illumination within

the cabinet was provided by an RB LED array (35 lmol m)2 sec)1;

MD Electronics; http://www.mdelectronics.co.uk).

DF images were collected immediately preceding lights off. The

cooled low light CCD camera was set in high scan mode with gain

set at low and binning set to 2 · 2. DF images were taken using a 1-

min exposure. The camera and lights were both automatically

controlled with WASABI imaging software (Hamamatsu Photonics;

http://www.hamamatsu.com). The images produced (RBF files)

were converted to TIFF files using WASABI. DF was quantified

using Metamorph (Molecular Devices Ltd; http://www.molecular

devices.com) to measure integrated luminescence for specificregions within an image. Background intensities, for each image,

were calculated and subtracted, to give a final DF measurement.

Measurement of the spectral emission and kinetics for DF

Arabidopsis seedlings were grown on MS media for 9 days. To

measure the spectral emission, seedlings were placed in the

imaging system and exposed to RB light of approximately

35 lmol m)2 sec)1 light for 1 min, and then the LED lights toggled

off. A 1-min exposure using the low-light imaging system described

above was taken through a vari-spec Liquid Crystal Tunable Filter

(CRI Inc.; http://www.cri-inc.com) tuned to 400 nm. The process was

Figure 5. DF and luciferase luminescence can be

assayed simultaneously in the plant.

Arabidopsis Ws seedlings containing CAB2:LUC

were grown on MS media containing 3% sucrose

and entrained under 12-h light/12-h dark cycles

for 9 days at 22°C before transferring to constant

RB light (35 lmol m)2 sec)1). DF and luciferase

luminescence were measured with a 2-h timeresolution for 96 h.

(a, b) The plots represent normalized averages

for DF and luciferase luminescence of 16 groups

of seedlings. (a) Ws CAB2:LUC n = 16 (filled

squares); (b) DF n = 16 (empty squares). Error

bars indicate SE.

(c) Period estimates plotted against their RAE.

WsDF n = 16(empty squares) andWs CAB2:LUC

n = 16 (filled squares).





then repeated, each time incrementing the filter by 20 nm from

400–720 nm.

For measuring the kineticsof DF, 9-day-old seedlings wereplaced

in the imaging system described above under RB light

(35 lmol m)2 sec)1). After 10 min the lights were turned off and a

series of five 10-sec exposures were taken.

DF rhythm analysis

The Arabidopsis plants were grown in groups of between 15–20

seedlings in 12-h light/12-h dark cycles at 22°C for 9 days. On dawn

of the 9th day the plants were placed in the imaging system at 22°C

in constant RB light. DF images were collected every hour as de-

scribedabove.The image acquisition andswitching of theLED array

was fully automated using the time-lapse function in WASABI. A

similar protocol was used for C. bursa-pastoris and lettuce seed-

lings. For maize, barley and K. fedtschenkoi , soil grown plants were

entrained in 12-h light/12-h dark cycles. For K. fedtschenkoi leaf

pairs 5, 6, 7 or 8 were used for imaging DF, as these leaf pairs

perform robust CAM CO2 fixation rhythms. Prior to the start of the

experiments, leaves were excised and placed on MS 1.5% agar

plates. For maize, leaves were excised and the cut end embedded in

MS media in test tubes. The test tubes were sealed with micro-pore

tape (Figure S8). For barley, leaves were cut into 1-cm pieces and

floated on SDW.

The DF images were processed as described above. The lumi-

nescence was normalized by subtracting the Y value of the best

straight line from the raw Y value. BRASS (available from http://

www.amillar.org) was used to carry out fast Fourier transformed

non-linear least-square analysis (Plautz et al., 1997) on each DF time

course series to generate period estimates and RAE.

Dual measurement of CAB2:LUC promoter activity and DF

Arabidopsis plants transformed with the CAB2:LUC reporter

construct were grown and entrained as described above. At 24 hprior to the start of the assay the plants were sprayed with 5 mM

luciferin (Biosynth; http://www.biosynth.com). The plants were

placed in the imaging system on dawn of the 9th day. After 95 min

the LED array was switched off and immediately a 1-min exposure

acquired. The system paused for 1 min in the dark, then a further

three, 1-min images were collected. The average signal of the final

three 1-min images was calculated and subtracted from the first

image, thus giving a value for the DF with luciferase luminescence

subtracted. Finally, the luciferase activity was measured by taking a

20-min image. The LED array was switched back on and the cycle

repeated for 96 h. The DF and luminescence data was processed as

described above and rhythmicity and periodicity scored using

BRASS.

ACKNOWLEDGEMENTS

The idea was originally conceived by J.H and A.H. Subsequent

experiments were designed and performed by A.H and P.D.G. The

C. bursa-pastoris experiments were performed by P.D and C.H, the

lettuce experiments by J.K, and barley experiments by R.S. We

would like to thank Dr Giles Johnson, University of Manchester

for critical reading of the manuscript and offering useful advice.

Research at Liverpool was funded by BBSRC grant BBS/B/11125

and Royal Society Grant R4917/1 to A.H. Funding for P.D and C.H

was provided by the Nuffield Foundation Science Bursaries

Scheme. Funding for J.K was provided by the Marie Curie Ac-

tions-Host fellowships for Early Stage Research Training (EST)

MEST-CT-2005-020526. Funding for R.S was provided by Libyan

Government.

SUPPORTING INFORMATION

Additional Supporting Information may be found in the onlineversion of this article:

Figure S1. The DF responseis not caused by LEDauto-fluorescence.

Figure S2. DF can be measured in both single Arabidopsis seed-

lings and excised Arabidopsis leaves.

Figure S3. The clock regulates the amount of DF and not the rate of

decay.

Figure S4. Arabidopsis circadian clock mutations affect DF rhythms.

Figure S5. DF rhythms can be measured in a range of plant species.

Figure S6. Robust DF oscillations can be visualized as non-uniform

waves of DF tracking across the leaves of K. fedtschenkoi .

Figure S7. Robust DF oscillations can be visualized as non-uniform

waves of DF tracking across the leaves of maize.

Figure S8. Picture of excised maize leaves imaged in the DF

experiment.

Please note: Wiley-Blackwell are not responsible for the content orfunctionality of any supporting materials supplied by the authors.

Any queries (other than missing material) should be directed to the

corresponding author for the article.

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