Phycologia (2007) Volume 46 (5), 503–512
Published 4 September 2007
Characterising nutrient-induced fluorescence transients (NIFTs) in nitrogen-stressed
Chlorella emersonii (Chlorophyta)
KIRSTEN SHELLY1*, TARA HIGGINS1,2, JOHN BEARDALL1, BAYDEN WOOD3, DON MCNAUGHTON3
AND
PHILIP HERAUD1
1
School of Biological Sciences and Centre for Biospectroscopy, Monash University, Clayton, Victoria, Australia
2
Department of Microbiology, National University of Ireland, Galway, Ireland
3
School of Chemistry and Centre for Biospectroscopy, Monash University, Clayton, Victoria, Australia
K. SHELLY, T. HIGGINS, J. BEARDALL, B. WOOD, D. MCNAUGHTON AND P. HERAUD. 2007. Characterising nutrientinduced fluorescence transients (NIFTs) in nitrogen-stressed Chlorella emersonii (Chlorophyta). Phycologia 46: 503–512.
DOI: 10.2216/06-55.1
Although determining the nutrient status of algae is highly desirable for water quality management, conventional
methods for assessing nutrient limitation are confounded by a range of theoretical and practical limitations. The current
paper examines the effectiveness of a novel rapid fluorometric technique, based on observations of nutrient-induced
transients in chl a fluorescence (NIFTs), in providing accurate measurements of algal nutrient limitation. The progress
of N-starvation in batch cultures of the freshwater chlorophyte Chlorella emersonii was followed and the NIFT
responses of cells to N-starvation and additions of both NH4+ and NO32 were characterised using a number of
techniques including NIFTs, conventional pigment analysis using UV/Visible spectrophotometry and Fourier transform
infrared (FT-IR) and Raman spectroscopy. Results showed that the addition of NH4+ and NO32 elicited distinct
changes in in vivo chl a fluorescence in N-limited Chlorella cells. The nature of the fluorescence change was dependent on
the form of nitrogen supplied, with NH4+ additions producing a more complex effect on fluorescence than NO32
additions. Interestingly, the greatest fluorescence response following NH4+ injection occurred prior to cells becoming
extremely N-starved. Despite Chlorella exhibiting many of the characteristic features of N-deficient cells such as lowered
capacity for photosynthetic electron transport (rETRmax), reduced Chl a : b–carotene ratios and increased levels of
carbohydrate accumulation relative to protein, values for the maximum effective quantum yield of Photosystem II
(WPSIIe-max) remained high over the course of N-starvation.
KEY WORDS: Chlorella emersonii, Nitrogen limitation, Nutrient-induced fluorescence transient (NIFT), Chlorophyll
fluorescence
INTRODUCTION
Nutrient availability in both marine and freshwater systems
frequently limits aquatic primary productivity, the most
common of limiting nutrients being nitrogen and phosphorus (Schindler 1977; Wynne & Berman 1980; Lean & Pick
1981). The ability to rapidly identify nutrient limitation,
and which nutrient in particular is limiting, would be of
great importance, both in understanding aquatic ecology
and for guiding water quality policies and management
practices. Historically, a number of techniques have been
used to determine the physiological health of algae, notably
nutrient-enrichment bioassays, and external and cellular
nutrient ratio analyses (as discussed by Hecky & Kilham
1988; Beardall et al. 2001b). However, most of these
techniques are not only complicated and time consuming,
they are also subject to potential artifacts, frequently
indicating potential limitation (Liebig limitation) in the
absence of other limiting factors rather than the in situ
nutrient status (Elser et al. 1990). As a result, the
interpretation and management of nutrient limitation in
aquatic systems using conventional techniques is oftentimes
difficult (Beardall et al. 2001b).
* Corresponding author (Kirsten.shelly@sci.monash.edu.au).
Nutrient limitation often is diagnosed in field populations of algae by chlorophyll a (Chl a) fluorescence (Kolber
et al. 1994). In particular, the chlorophyll fluorescence
parameter WPSIIe-max (the maximum quantum yield of PS
II; 5 Fv/Fm) has been used as an index of microalgal
physiological status. Recent studies spanning marine and
freshwater organisms, involving both field assemblages and
laboratory cultures of algae, have indicated that changes in
steady state chlorophyll fluorescence and WPSIIe-max
following nutrient addition could potentially serve as
a diagnostic tool for assessing nutrient limitation in
microalgae. Turpin & Weger (1988) originally reported
that, if phosphorus or nitrogen is added to phytoplankton
cultures limited in that particular nutrient, a rapid but
transient change (within minutes) in fluorescence is seen.
This change in chlorophyll fluorescence has been termed
a nutrient-induced fluorescence transient, or NIFT (Wood
& Oliver 1995). These fluorescence transients have been
reported for the resupply of carbon (Miller et al. 1991),
nitrogen (Holmes et al. 1989; Wood & Oliver 1995; Young
& Beardall 2003b) and phosphorus (Wood & Oliver 1995;
Gauthier & Turpin 1997; Roberts 1998; Beardall et al.
2001b) to suitably deficient cells. There is also some
evidence that this phenomenon occurs with silicate (Lippemeier et al. 2001). The fluorescence response generally is
not observed if a nonlimiting nutrient or distilled water is
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added to the cells (Beardall et al. 2001a, b; Holland et al.
2004). Although the precise physiology behind NIFTs is
still poorly understood, it is believed that the transient
change in fluorescence is attributable to a reallocation of
energy from photosynthesis to nutrient uptake.
Chlorophyll a and b-carotene content, as well as the
macromolecular composition of algae, also change with
changing cell nutrient status. Raman spectroscopy and
Fourier transform infrared (FT-IR) methodologies are
novel ways in which to determine both pigments, such as
chlorophyll a and b-carotene, and macromolecular content
of cells, respectively. FT-IR spectroscopy previously has
been used to carry out rapid, simple, concurrent and
noninvasive analysis of cellular components. Giordano et
al. (2001), for example, showed that FT-IR spectroscopy is
a reliable way to precisely determine the relative concentration of silica, protein, carbohydrates and lipids in the
diatom Chaetoceros muelleri. Given its ability to minimise
disturbance of the intracellular environment due to reduced
manipulation of the sample, FT-IR spectroscopy appears to
be a powerful analytical tool. Previous methods for
measuring cellular composition such as wet chemistry for
internal pools of macromolecules, or estimates of changes
in elemental ratios (e.g. C : N and N : P), are not
necessarily specific to the limiting nutrient, and often are
prone to interference from debris in natural assemblages
(Beardall et al. 2001a). Cell protein profiles run on gel
electrophoresis separate out proteins, which can identify
changes in levels of specific proteins, but the sample sizes
needed are often large and the method is time consuming
(Beardall et al. 2001a). FT-IR spectroscopy takes around
20 seconds to acquire a spectrum, and the sample size is
small enough not to perturb the culture system (,4 ml
culture is needed), whilst still allowing accurate assessment
of the changes in macromolecular content of the cell.
Raman spectroscopy can be used with live cells (Wood et al.
2005) and allows spectra to be taken on a single cell basis
(Heraud et al. 2006; Heraud et al. 2006). This technique is
an efficient and simple tool for making measurements of
chlorophyll a and b-carotene, rather than simply measuring
carotenoids in bulk samples using conventional spectroscopic methods.
To date, the exploration of NIFTs as a reliable index of
nutrient limitation has been restricted to a small number
of algal species. Few investigations have characterized
these NIFT responses thoroughly (but see the studies on
the marine chlorophyte Dunaliella tertiolecta by Young &
Beardall 2003a, b). Holland et al. (2004) also reported
briefly on the occurrence of phosphate and ammoniuminduced NIFTs in Chlorella, as well as in the cyanobacterium Oscillatoria and mixed field assemblages. In order to
further explore the possibility of using this technique as an
indicator of nitrogen stress in algae, the current study
followed the progress of N-starvation in batch cultures of
the freshwater chlorophyte Chlorella emersonii and characterized in detail the NIFT response shown by this
organism to the respective addition of both NH4+ and
NO32, as well as investigating the changes in chlorophyll
a : b-carotene ratios and macromolecular composition of
the cells.
MATERIAL AND METHODS
Culture conditions
Chlorella emersonii was cultured under axenic batch growth
conditions at 18uC using 250 ml glass vessels, which were
exposed continuously to photosynthetically active radiation
(PAR) (120 mmol quanta m22 s21) provided by Sylvannia
Gro-lux WS (Osram Sylvania Ltd., Danvers, MA, US)
fluorescent tubes. The cultures were maintained in MS/2
medium (Morris & Syrett 1965), in which only the nitrogen
input was adjusted. Cultures were initiated with starting
KNO3 concentrations of 50 mM (‘low-N cultures’) or
100 mM (‘high-N cultures’) in order to allow some initial
growth, and 3 replicate 250-ml cultures were set up for both
KNO3 levels. All other nutrients were in large excess. The
two differing N treatments were used to induce starvation at
different rates in order to improve our ability to resolve
when cells start to show the onset of the NIFT perturbation.
Measurement of WPSIIe-max
Subsamples were withdrawn regularly from cultures to take
chlorophyll fluorescence measurements. These were carried
out with a PhytoPAM (Walz, Effeltrich, Germany), which
was modified such that the cuvette could be maintained at
the growth temperature and stirred from above with
a motorised stirring unit. Prior to acquisition of the
PhytoPAM measurements, cells were dark adapted in the
cuvette for 15 min. WPSIIe-max then was measured after the
application of a saturating pulse and calculated according
to Van Kooten & Snel (1990). Rapid light curve (RLC)
measurements were carried out on the PhytoPAM modified
as above. The actinic light was altered after each
measurement from 1 to 295 mmol quanta m22 s21, with
each light intensity being applied for 30 seconds. The
maximum relative electron transport rate (rETRmax) was
calculated by fitting the rETR and irradiance data to the
equations of Eilers & Peeters (1988).
CHLOROPHYLLS AND CAROTENOIDS: Appropriate, duplicate
volumes of sample (between 5 and 20 ml, depending on the
cell density of the culture) were filtered through a GF/C
filter. Filters were wrapped in aluminium foil and frozen for
later processing. Frozen filters were extracted in 10-ml
centrifuge tubes using 5 ml of a 1 : 1 v/v mixture of
dimethyl sulfoxide (DMSO) and 90% acetone (Burnison
1980). Tubes were well shaken to accelerate pigment
extraction and placed in darkness in a refrigerator.
Following overnight extraction, tubes were centrifuged
and the supernatant collected. Sample absorbances were
read on a Cary 50 spectrophotometer and chlorophyll and
carotenoid concentrations were calculated according to
Golterman & Clymo (1971).
FT-IR AND RAMAN SPECTROSCOPY: FT-IR spectra of dried
cell pastes were collected on a Bruker IFS55 EQUINOX
FT-IR spectrometer equipped with a liquid nitrogen cooled
HgCdTe and using a Golden Gate (TM Specac, Woodstock, GA, US) single bounce diamond attenuated total
reflectance (ATR) accessory. Moist cell pastes (approx 5 3
109 cells/ml) were air-dried onto the diamond window of the
Shelly et al.: Nitrogen limited NIFTs
505
FT-IR accessory before spectra were collected. Typically,
50 scans were taken with a resolution of 6 cm21. Raman
spectra of single viable algal cells were recorded on
a Renishaw system 2000 (equipped with a Peltier cooled
CCD detector) using a 782 nm excitation line from a diode
laser and a modified BH2-UMA Olympus optical microscope with a Zeiss 3 60 water immersion objective running
under Renishaw-WiRETM 2 software. Ten microliters of
packed cells were added to a 6 cm diameter aluminium
sputter coated Petri dish that was further coated in 0.01%
poly-l-lysine to affix the cells to the bottom of the Petri dish.
Power at the sample was ,2–3 mW for a 1–2 mm laser spot
size. Spectra were recorded between 1800–200 cm21 with
a resolution of ,1–2 cm21. For each spectrum 1 scan was
accumulated and the laser exposure for each scan was
10 seconds for a population of 30 individual cells under
each condition. The spectra were then baseline corrected
using the rubber band method of the OPUSTM spectroscopic software (Bruker, Germany), corrected using the
multiplicative scattering correction within The UnscramblerTM (Camo, Norway) and averaged.
Nutrient-induced fluorescence transients
NIFTs (Beardall et al. 2001a) were investigated during the
course of N-starvation following the method of Holland et
al. (2004). A 3-ml sample was taken from each of the batch
cultures and placed in a mirrored glass fluorometer cuvette
with a magnetic stirrer. The sample was placed in a Hitachi
2000 spectrofluorometer (EX at 435 nm and EM at
680 nm) and steady state fluorescence (Ft) values were
measured until stable. Once fluorescence had stabilised,
a 30-ml sample of either NH4+ or NO32 solution was
injected into the cuvette, to give a final concentration of
100 mM. The observed changes in Ft then were recorded
and the characteristic features of fluorescence changes
following the injection were quantified (see Fig. 1).
Subsamples also were cross-checked and no changes in
fluorescence, at any point during starvation, were evident
upon additions of either P (10 mM PO432) or dH2O,
indicating that nitrogen was the limiting nutrient. Unless
otherwise stated, differences between parameter values were
analysed statistically by students t test.
RESULTS
WPSIIe-max
There was no significant change (,5%; P . 0.05) in
WPSIIe-max over the 10-day experiment (Fig. 2a). Little
variation was observed between batch cultures of different
starting N concentrations, with both ‘low-N’ and ‘high-N’
treatments dropping to a similar WPSIIe-max value within 10
days of starvation (WPSIIe-max 0.740 and 0.743, respectively).
CHLOROPHYLL AND CAROTENOIDS: Cellular chlorophyll
concentrations decreased dramatically as N-starvation
progressed (Fig. 2b). Cultures initiated at both ‘low-N’
and ‘high-N’ levels showed a similar pattern, with
Fig. 1. An example of the characteristic features of (a) a NO32–
nutrient-induced fluorescence transients (NIFTs) and (b) a NH4+–
NIFT in ‘low-N’ batch cultures of Chlorella emersonii.
chlorophyll dropping over fivefold, from 397.0 and
428.2 pg/cell on day 1 to 79.9 and 75.9 pg/cell on day 10
in ‘low-N’ and ‘high-N’ cultures, respectively. Carotenoid : Chlorophyll ratios (Fig. 2b) increased substantially
as the cells became more N-starved, from 1.19 and 0.71 on
day 1, to 3.24 and 3.01 on day 10, in the ‘low-N’ and ‘highN’ cultures, respectively. Values for chlorophyll content
and carotenoid : chl ratio were not significantly different (P
. 0.05) between ‘low-N’ and ‘high-N’ cultures except on
day 4, when the chlorophyll content was lower in the ‘lowN’ cells (P , 0.05).
RLC MEASUREMENTS: Measurements of rETR as a function of photon flux (Fig. 2c) indicated that, as N-starvation
advanced, light saturated rates (rETRmax) declined by 27%
and 32% in the ‘low-N’ and ‘high-N’ cultures, respectively
(P . 0.05). The ‘low-N’ batch cultures exhibited 10–16%
lower rETRmax compared with cells grown initially at ‘highN’ (P . 0.05).
NIFT CHARACTERISTIC RESPONSE: Fig. 1 shows the characteristic NIFT response following NO32 and NH4+ additions
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Fig. 2. (a) Changes in photosynthetic parameters and pigments of Chlorella emersonii batch cultures following the transfer of cells to ‘low-N’
(black bars, filled symbols) and ‘high-N’ (open bars, open symbols) batch cultures. (a) PSIIe-max; (b) total chlorophyll (bars) and
carotenoid : chlorophyll ratio (lines); (c) light saturated rates of relative electron transfer (rETRmax). All data shown are the mean of
replicate cultures, n 5 3, with the exception of chlorophyll/carotenoid data where n 5 6. Error bars, where shown, give standard deviations.
to batch cultures of Chlorella emersonni as N-starvation
progressed, normalised to pre-injection steady-state fluorescence (Ft). It is apparent that NH4+ additions (Fig. 1b)
had a more complex effect on chl a fluorescence than did
NO32 (Fig. 1a). Following injection of NO32, there was
a slow, steady decline in fluorescence after 30–300 seconds
(DDrop), followed by a very slow, gradual increase in
fluorescence emission over a longer time interval, to ,75%
of initial, pre-injection fluorescence (Ft) after 400 seconds.
In contrast, NH4+ injection caused an initial small but
marked (,10%) rise in fluorescence output after 10–
50 seconds (DPeak), quickly followed by a rapid drop in
fluorescence after 50–100 seconds (DDrop), with fluorescence reaching a minimum (90%) before returning to a new,
lower steady state value (91–94% of Ft) after 130 seconds
(Fig. 1b). Addition of a nonlimiting nutrient (e.g. PO432) or
control (dH2O) to the ‘low-N’ or ‘high-N’ Chlorella cultures
produced no measurable change in fluorescence output, nor
did the addition of NH4+ or NO32 to Chlorella cells grown
in nutrient replete medium (data not shown).
TIMING OF THE ONSET OF NIFTS: In the low-N Chlorella
cultures, the addition of NO32 on day 1 produced no
noticeable change in fluorescence (Fig. 3b), although the
addition of NH4+ elicited a characteristic NH4+ NIFT,
which was small in magnitude (,5% DPeak, ,5% Ddrop;
Fig. 3a). By day 4, a more developed NIFT was evident and
at its most prominent for both NH4+ (32% change in Ft,)
and NO32 (23% change in Ft,) additions in the ‘low-N’
cultures, before beginning to disappear at day 7. By day 10,
a NO32 NIFT had almost completely disappeared in the
‘low-N’ cultures (5% change in Ft; Fig. 3b), although the
NH4+ NIFT was still evident, although of a considerably
smaller magnitude than on day 4 (9% change in Ft;
Fig. 3a).
The % D Drop and % D Peak following NH4+ addition
are quantified in Figs 1a, 4b, respectively. Both ‘low-N’ and
‘high-N’ batch cultures showed a significant (P , 0.05) rise
to maximum peak in fluorescence following NH4+ injection
on day 4 (18 and 28%; Fig. 4a). The % DDrop following the
injection of NH4+ also increased significantly (P , 0.05) on
day 4, before rapidly dropping as N-starvation progressed
further (Fig. 4b). There were no significant differences in
this pattern between ‘low-N’ and ‘high-N’ batch cultures
(P . 0.05). By day 10, the % DDrop in response to NH4+
injection had declined significantly (P , 0.05) in both the
‘low-N’ and ‘high-N’ cultures (Fig. 4a), while the % Dpeak
remained high with a significant difference (P , 0.05)
remaining on day 10 between ‘low-N’ and ‘high-N’ cultures
in the ‘high-N’ samples (19%; Fig. 4b).
NIFTs showed a less distinct pattern in the % Ddrop
following NO32 injection (Fig. 4c), compared with NH4+
injection (Fig. 4b). Both ‘low-N’ and ‘high-N’ cultures
showed a large increase in the % Ddrop on day 4 following
NO32 addition (31 and 30%, respectively). However, by
day 7, ‘low-N’ batch cultures showed a decline in % Ddrop
in response to NO32 addition, whilst the % Ddrop in the
‘high-N’ batch cultures remained significantly higher (e.g.
19% on day 10) compared to ‘low-N’ cultures (10% on day
10; P , 0.05; Fig. 4c).
There appeared to be little correlation between the onset
of NIFTs and trends in WPSIIe-max for Chlorella which, as
already stated, remained consistently high over the course
of the experiment (Fig. 2a). Indeed, the cultures exhibited
the most marked NIFT responses on day 4, when WPSIIemax values were relatively high (0.76 and 0.77 for the lowand high-N cultures, respectively).
FOURIER TRANSFORM INFRARED SPECTROSCOPY: FT-IR
spectra were acquired for both ‘low-N’ and ‘high-N’ batch
cultures. Amide I and II bands arising from protein and
a broad band associated with the C-O stretches of
carbohydrate were identified in the regions from 1630–
1677 cm21, 1520–1560 cm21 and 990–1100 cm21, respectively (Fig. 5A). Fig. 6a shows clearly that, as N-starvation
progressed, both the carbohydrate : amide I and carbohydrate : amide II ratios increased substantially in both ‘highN’ and ‘low-N’ starved cultures, and were shown to be
significantly different (P , 0.05, analysis of variance) from
the replete (day 1) cells. Principal component analysis also
was performed, which supported the discrete difference
between day 1 and day 10 data for both ‘high’ and ‘low’ N
Shelly et al.: Nitrogen limited NIFTs
507
Fig. 3. (a) Graph showing nutrient-induced fluorescence transients (NIFTs) following NH4+ injection (represented by the vertical line) on
days 1 (closed circles), 4 (open circles), 7 (closed triangles) and 10 (open triangles). (b) Graph showing NIFTs following NO3 injection
(represented by the vertical line) on days 1 (closed circles), 4 (open circles), 7 (closed triangles) and 10 (open triangles). All data is for the
‘low-N’ cultures and are normalised to steady state values prior to N-addition. Typical traces have been shown in this graph and displaced
by 10% for clarity; any variation between traces can be seen in Fig. 4. The line on graph 4a indicates the percentage change in fluorescence.
cells. A loading plot is used to assign coefficients to relevant
PCs from an x, y-variable data set. Loadings express
relations between variables (i.e. wavenumber values), and
enable one to tell which variables are dominating or
influencing the model and how they are associated with one
another. Analysis of the PC1 loadings plot in this study
confirmed that the separation along PC1 primarily was
associated with changes in the intensity of carbohydrate
bands (data not shown).
RAMAN SPECTROSCOPY: Raman spectroscopy was performed on ‘low-N’ and ‘high-N’ cultures. Chl a levels
(identified as the integrated area between 1314 to
1337 cm21, Fig. 5B) showed a decline in the ‘low-N’
cultures after a short period of starvation and b-carotene
(identified as integrated band areas from 1132–1172 and
1491–1518 cm21, referred to as b-carotene bands 1 and 2,
respectively; Fig. 5B; see Wood et al. 2005 for a full
description of band assignments). A clear decline over time
in the chl a : b-carotene ratio is evident in both N treatments
(Fig. 6b). All changes in cells from day 1 (replete) to day 10
(‘high’ and ‘low’ N) were significantly different (P , 0.05,
analysis of variance). Principal component analysis also was
performed (data not shown), which supported the discrete
differences between day 1 and day 10 data for both ‘high’
and ‘low’ N cells. Loadings plots, in this case such
differences were principally influenced by the changes in
the b-carotene bands 1 and 2 (data not shown).
Fig. 4. (a) Percentage difference in DPeak following NH4+ injection into ‘low-N’ (dark bars) and ‘high-N’ (light bars) cultures of Chlorella
emersonii cells. (b) Percentage difference in DDrop following NH4+ injection into ‘low-N’ (dark bars) and ‘high-N’ (light bars) cultures of C.
emersonii cells. (c) Percentage difference in DDrop following KNO3 injection into (‘low-’ and ‘high-N’) cultures of C. emersonii cells. All
data shown are the mean of replicate batch cultures, n 5 3. Error bars represent the standard deviations.
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Phycologia, Vol. 46 (5), 2007
Fig. 5. Representative curves from Chlorella cells on days 1 (replete) and 10 (‘high-’ and ‘low-N’) from (A) Fourier transform infrared
(FT-IR) data showing the amide I (1630–1677 cm21), II (1520–1560 cm21) and carbohydrate (990–1100 cm21) bands. (B) Raman
spectroscopy data showing bands for b-carotene (1491–1548 cm21) and chl a (1314–1327 cm21).
DISCUSSION
Nitrogen limitation can affect a number of processes in
phytoplankton including photosynthesis, photochemical
energy conversion and protein synthesis (Berges et al.
1996; Jansen et al. 1996; Masi & Melis 1997). This study
showed that cellular Chl a : b-carotene ratios, measured
using both a conventional spectroscopic assay as well as
Raman spectroscopy, declined in the freshwater chloro-
phyte Chlorella emersonii as N-starvation progressed.
Raman spectroscopy can be an informative tool for the
study of many biological systems because of the large
number of active vibrational modes and the relative lack of
spectral interference from water. The strong enhancement
of many b-carotene bands at 785 nm makes Raman an
excellent diagnostic tool to gain quick and precise information regarding chl a and b-carotene content (Wood
et al. 2005). It is far more specific than conventional
Fig. 6. Integrated areas for Chlorella cells on days 1 (replete) and 10 (‘high-’ and ‘low-N’) showing (a) Fourier transform infrared (FT-IR)
data for ratios of carbohydrates: amide I (dark bars) and amide II (light bars). (b) Raman data showing ratios of chl a : b-carotene (1314–
1327 cm21 and 1491–1548 cm21 respectively). All data for both FT-IR and Raman on day 10 (‘high-’ and ‘low-N’) were shown to be
significantly different from day 1 (replete; P , 0.05, analysis of variance) marked with an asterisk (*).
Shelly et al.: Nitrogen limited NIFTs
spectroscopic methods (e.g. Golterman & Clymo 1971),
which only measure carotenoids as a group of compounds,
rather than specifically b-carotene, and is much faster than
high performance liquid chromatography (HPLC), which,
while giving information on specific pigments, can be time
consuming. Raman spectroscopy also can be applied to
single cells rather than the bulk populations required for
conventional spectrophotometric and HPLC approaches.
It is well established that N-deficiency in eukaryotic algae
can lead to decreases in antenna size and a decrease in the
chl a : carotenoid ratio (Turpin 1991).This decline in chl a,
as well as in proteins (see discussion of FT-IR data below),
appears to imply either that the photosynthetic complex of
Chlorella was compromised, or simply that there were
reduced levels of photosynthetic units per cell which
remained fully functional and were capable of maintaining
photosynthetic efficiency as nitrogen starvation progressed.
Photosynthesis can be affected by nitrogen limitation
directly through the reduction in energy-collecting efficiency brought about by the loss of chl a and the concomitant
increase in carotenoid pigments which are photochemically
inactive (Herzig & Falkowski 1989; Berges et al. 1996).
Photochemical energy conversion is affected by a reduction
in protein synthesis, which ultimately affects the protein
content of the PSI and PSII reaction centres. Although
prior research has found that PSII content is affected more
by nitrogen limitation than is PSI, it must be acknowledged
that this may vary between species (Behrenfeld et al. 1996;
Berges et al. 1996). Given that the PSII proteins, such as
D1, have a rapid turnover, a decline in nitrogen availability
could (1) reduce the supply of amino acids for repair; and
(2) through effects on electron transport, reduce the
adenosine 59-triphosphate (ATP) available to drive
(re)synthesis of this protein, in effect enhancing the decline
in PSII photosynthesis.
In the current study, however, Chlorella showed no
marked change in WPSIIe-max over the course of starvation.
This observation was surprising, because the photosynthetic capacity of algal cells (including WPSIIe-max) typically
declines markedly during N-limitation (Falkowski et al.
1989; Geider et al. 1993; Shelly et al. 2002; Young &
Beardall 2003a). The fact that little difference in WPSIIe-max
was observed between both the ‘low-N’ and ‘high-N’
cultures also suggested that the degree of N-starvation
had little impact on photosynthetic capacity in Chlorella,
with likely implications for the NIFT observations in the
two treatments. In N-stressed batch cultures of Dunaliella,
for example, WPSIIe-max decreased from more than 0.6 to
0.2 over 4 days (Young & Beardall 2003b). Other studies,
however, have reported a contrasting trend similar to that
observed here for Chlorella, in which WPSIIe-max values
have remained consistently high over the course of nutrient
starvation (Cullen et al. 1992; MacIntyre et al. 1997;
Parkhill et al. 2001). Equally high WPSIIe-max values also
were observed during the current study in C. emersonii
grown in N-limited semicontinuous cultures (Shelly et al.,
unpublished data). Viewed collectively, these findings
appear to suggest that WPSIIe-max responses to nutrient
stress in algae are species-specific and that WPSIIe-max is not
a robust indicator of nutrient stress in all algae (or, at least,
not in N-limited Chlorella; see Kruskopf & Flynn 2006 for
509
a critique). One hypothesis is that algal cells can use ATP
produced from cyclic electron flow around PSI, in order to
repair damage to PSII or to drive nutrient uptake through
ATP-dependent pumps (Berges 1997). Whether this mechanism would be capable of maintaining the integrity of PSII
and sustaining the high WPSIIe-max observed here for Nstarved Chlorella is not known.
FT-IR spectroscopy provides information on vibrationally active functional groups associated with macromolecules in biological samples, reflecting both the molecular
structure and environment of the sample. The FT-IR
technique has been used to examine isolated macromolecules
such as nucleic acids, proteins, lipids and polysaccharides,
with some studies having extended this work to investigate
whole organisms, such as marine microalgae (Giordano et
al. 2001) and higher plants (Stuart 1997; Heraud et al. 2006).
Previous studies (Parker 1971) on the macromolecular
content of cells have allowed a summary of band assignments to be collated. These studies also have validated the
bands with standards, allowing the characterisation of
amide I and amide II (proteins) by two strong bands at
,1650 cm21 and ,1540 cm21, assigned to specific vibrations of the peptide moiety. Carbohydrates show characteristic C-O bonds at ,1033, 1080 and 1150 cm21, while the
carbonyl group is observed at 1740 cm21 (Kansiz et al.
1999). FT-IR analysis in the current study clearly revealed
dramatic spectral differences between N-replete (day 1) and
N-limited (day 10) Chlorella cultures. As N- starvation was
induced, there was a shift towards carbohydrate accumulation, relative to protein. It is well established that N-limited
algal cells generally accumulate starch or lipid and use this
as a source of carbon for amino acid synthesis (Turpin
1991). Documentation of protein loss in algal cells during Nlimitation is extensive (Richards & Thurston 1980; Kolber et
al. 1988; La Roche et al. 1993; Quigg and Beardall 2003;
Young and Beardall 2003b). Smith et al. (1989) showed that
allocation of ATP to protein synthesis declined when Nlimitation was imposed on the ice alga Nitzschia seriata,
although allocation towards lipids increased. With certain
exceptions (MacIntyre et al. 1997), studies have shown large
declines in the pools of Rubisco under nitrogen limitation
(Beardall et al. 1991). Young & Beardall (2003a) suggested
that, under some conditions, the Rubisco protein can act as
a store of N, which then can be mobilised to enable
maintenance of other N-requiring functions, such as
photosynthetic electron transport, under stress conditions.
Given that Chlorella exhibited little change in WPSIIe-max
during the current experiment, we can only assume that the
turnover rate of the D1 protein remained intact and that the
shift in cellular energy allocation from protein towards
carbohydrates synthesis had little effect on the photosynthetic apparatus of Chlorella.
As starvation progressed, the addition of NH4+ and
NO32 to the N-limited Chlorella cells elicited characteristic,
transient changes in in vivo chl a fluorescence (NIFTs). The
configuration of the NIFT response was shown to depend
on the form in which the nitrogen was supplied: NH4+
elicited a small, sudden rise in fluorescence, which then
dropped sharply, before recovering to a new, lower steady
state; NO32 additions, in contrast, caused a much more
gradual, steady decline in fluorescence. Interestingly, the
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Phycologia, Vol. 46 (5), 2007
latter response differed from that found in Dunalliella
tertiolecta by Young & Beardall (2003b), supporting predictions that the precise nature of the fluorescence change
depends on the particular algal species (Wood & Oliver 1995;
Holland et al. 2004). In contrast, the NH4+–induced
responses observed here were very similar to those reported
by Holland et al. (2004) for N-limited Chlorella and by
Young & Beardall (2003b) for N-starved D. tertiolecta,
comprising an initial rapid and distinct peak in fluorescence,
followed by a trough and then a gradual recovery in
fluorescence to a new, lower, steady state level. As in previous
studies (Beardall et al. 2001a, b; Young & Beardall 2003b),
fluorescence perturbations were not seen in N-replete
Chlorella cultures following additions of NH4+ and NO32,
or in N-limited cultures following the addition of a nonlimiting nutrient (e.g. PO432) or control i.e. water (data not
shown). It is worth pointing out that we investigated the
responses of NH4+ and NO32 addition to cells that previously
had been grown on nitrate. Whether these responses differ for
cells grown on ammonia remains to be seen.
Results from the current study on N-limited Chlorella
indicated that the timing of NIFTs (i.e. their onset and
disappearance) varied according to both the starting
nitrogen concentration in the medium and the form of
nitrogen (NH4+ or NO32) supplied. For example, ‘high-N’
cultures resupplied with NO32 showed a greater % Ddrop
for an extended period of time compared with ‘low-N’
cultures, presumably reflecting the delayed starvation of the
former cultures, in which cells were slower to reach the
critical stage at which they lost the ability to show a NIFT.
In ‘low-N’ cultures (initially 50 mm KNO3), NH4+ –NIFTs
were, however, present from day 1 through to day 10; this is
a longer timeframe than that reported by Holland et al.
(2004), who observed NH4+–NIFTs over a period of 4 days
in N-starved Chlorella cultures. In contrast, NO32–NIFTs
appeared to have a smaller window of occurrence, being
first present on day 4 and having completely disappeared by
day 10. Young & Beardall (2003b) similarly recorded the
persistence of NH4+–induced fluorescence perturbations for
several days after additions of NO32 had ceased to elicit
a fluorescence response.
Both ‘low-N’ and ‘high-N’ cultures showed maximum
changes in fluorescence following either NH4+ or NO32
injections on day 4, indicating the greatest fluorescence
transients occurred prior to cells becoming extremely Nstarved. Other studies likewise have shown that the NO32induced NIFT disappears under extreme nutrient limitation
(Roberts 1998; Young 1999). One explanation is that, in
order to respond to NO32, cells must possess an induced
and optimised NO32 uptake and/or reduction capacity,
mechanisms which are only active in cells currently or
recently supplied with external NO32 (Young & Beardall
2003b). Furthermore, recovery of NO32-uptake in Nstarved D. tertiolecta was shown to require protein
synthesis (Young 1999). After prolonged starvation, the
loss of fluorescence perturbations seems to suggest that the
NO32/NO22 flux into the chloroplast is inadequate to
induce the effect of N-assimilation on photosynthetic
suppression.
The fluorescence transients exhibited following NH4+ or
NO32 injection to N-limited Chlorella presumably are
induced by the uptake and assimilation of the limiting
nutrient, nitrogen. N-uptake and assimilation in algae is
dependent on photosynthesis for energy and carbon
skeletons, and involve metabolic processes in both chloroplasts and mitochondria (Turpin et al. 1997). NH4+ or
NO32 assimilation and uptake can influence cellular charge
equilibria and, therefore, vary the relative requirement for
ATP by photosynthesis and nitrogen metabolism (Turpin &
Weger 1988; Holmes et al. 1989). Weger & Turpin (1989)
showed that the decrease in NH4+ assimilation, in the
absence of mitochondrial electron transport chain activity
during photosynthesis, is not due to a limitation in ATP
supply but rather to a shortage of carbon skeletons for
amino acid synthesis. Holmes et al. (1989) also found that
resupply of NH4+ to N-limited Selenastrum minutum
resulted in a drop in Rubisco levels and limitation of
photosynthetic carbon fixation, which they postulated was
due to increased requirements for carbon skeletons in the
synthesis of amino acids. Studies on S. minutum, using the
electron transport inhibitor, DCMU, resulted in an 82%
decrease in the rate of NH4+ assimilation, although NO32
assimilation was unaffected (Weger & Turpin 1989). It
appears that, once the nutrient pulse has been added to the
limited cells, amino acid turnover is stimulated via
a reallocation in the source of these carbon skeletons from
the Calvin cycle to stored carbohydrates, such as starch
(Turpin & Weger 1988; Wood & Oliver 1995).
Ammonium addition to both high- and low-N batch
cultures resulted in an increase in DPeak within about
10 seconds; this is approximately the same timeframe that
Turpin & Weger (1988) found for a decline in O2 evolution
in S. minutum, signalling a suppression of photosynthesis as
energy was reallocated to N-uptake and assimilation.
DDecline followed the DPeak at around 15 seconds, which
also seems to correspond with the timeframe for completion
of ammonium assimilation and the recovery of photosynthesis and O2 evolution (Turpin & Weger 1988). Additions
of NO32 or NH4+ to nutrient-replete Chlorella cells elicited
no changes in chl a fluorescence. It is likely that high
glutamine/glutamate ratios in the chloroplasts of nutrient
replete cells maintain a priority for photosynthesis over N
assimilation (Flynn 1991), so that N additions do not lead
to changes in the relative use of ATP and electrons and so
do not cause perturbations in chl a fluorescence.
Young & Beardall (2003b) found that the resupply of
NH4+ and NO32 to N-stressed D. tertiolecta cells also
produced changes in nonphotochemical quenching, probably due to state transitions resulting from the reallocation
of light harvesting components between PSI and PSII
(Muller et al. 2001). For instance, S. minutum has been
shown to exhibit a reversible transition from state I to state
II under N-limitation, diminishing PSII activity and
elevating the energy accessible to PSI (Turpin 1991). This
would subsequently increase the ATP/nicotinamide adenine
dinucleotide phosphate (NADPH) production ratio, in line
with the metabolic requirements of NH4+ assimilation.
Assimilation of NO32 and NO22 has an ATP/NADPH
production ratio similar to that of CO2 assimilation, and
these ions were not found to induce state transitions in S.
minutum (Turpin 1991). State transitions could be a method
in which algae maintain equilibrium of ATP and NADPH
Shelly et al.: Nitrogen limited NIFTs
production with changing metabolic demands. Whether
a state transition is responsible for the fluorescence changes
observed in the present study is yet to be determined.
Unlike traditional mechanisms for assessing nutrient
limitation in algae, NIFTs constitute rapid changes in
chlorophyll fluorescence over time scales of minutes, thereby
potentially offering an immediate indication of the nutrient
status of a particular algal population. Whilst some
parameters were measured using pulse amplitude modulated
(PAM) fluorometers, steady state fluorescence for NIFT
experiments were measured using the more simplistic Hitachi
Spectrophotometer, paving the way for field experiments to
be carried out on simple, affordable fluorometers. Previous
studies assessing the applicability of the technique for
investigating in situ algal nutrient limitation have reported
encouraging results for nitrate-induced NIFTs in the green
algae Dunaliella tertiolecta (Young & Beardall 2003b) and
for ammonium and phosphate-induced NIFTs in Chlorella
emersonii and the cyanobacterium Oscillatoria sp. (Holland
et al. 2004), as well as in natural, mixed field assemblages
(Wood & Oliver 1995; Holland et al. 2004). The current
characterisation of the distinct fluorescence responses of Nstarved Chlorella emersonii to NH4+ and NO32 additions
provides further evidence of the potential of NIFTs both for
identifying N-limitation generally and for assessing the
relative importance to algal nutrient of different nitrogen
sources. The apparent insensitivity of the onset and
magnitude of fluorescence changes to the initial N concentrations (50 and 100 mM NO3 in the ‘low-N’ and ‘high-N’
cultures, respectively) may be a caveat to the precision
and application of the technique as a diagnostic tool for the
degree of nutrient limitation. The observed high WPSIIe-max
values, however, indicate that Chlorella emersonii cells
may behave atypically under N-limiting conditions by
apparently maintaining a high photosynthetic efficiency.
Thus, the interaction between N-assimilation and photosynthetic suppression believed responsible for causing the
fluorescence transients may likewise be atypical in Chlorella.
Additional work is needed in order to better elucidate the
physiology behind the NIFT and to assess the variability, or
otherwise, of nutrient-induced fluorescence perturbations in
different algal species under varying levels of nutrient
deficiency.
ACKNOWLEDGEMENTS
This work was supported by the Australian Research
Council. Tara Higgins was a visitor to Monash University on an Endeavour Fellowship awarded by the
Australian Department for Education, Science and Training (DEST).
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Received 14 July 2006; accepted 30 April 2007
Associate editor: Charles Amsler