J. Exp. Mar. Biol. Ecol., 1991, Vol. 146, pp. 181-191
181
Elsevier
JEMBE 01547
Determination of nitrate reductase activity in Ulva rigida
C. Agardh by the in situ method
A. Corzo and F . X . Niell
Department of Ecology, University of MMaga, Campus de Teatinos, MMaga, Spain
(Received 6 April 1990; revision received 7 August 1990; accepted 26 October 1990)
Abstract: Assay conditions for determining in situ nitrate reductase (NR) activity in Ulva rigida C. Agardh
were studied. The final composition of the incubation medium was: 0.1 M phosphate buffer, 0.5 mM EDTA,
0.1% l-propanol, 30 mM KNO3 and 10 #M glucose. The optimal pH was 8, which is greater than that
usually found in higher plants (7.5) and close to the pH of sea water. Samples were incubated in the dark
at 30 ° C. A short incubation time (30 min)was more satisfactory than longer ones for determination of the
initial rate, and to prevent any limitation of the NO3 reduction rate for the assay conditions themselves.
Surfactants such as Tergitol NP-10 and Triton X-100 were less effective than 0.1% l-propanol. Incubation
in anaerobic conditions is a critical point in the in situ NR assay in Ulva rigida. The two methods used;
vacuum infiltration for 10 min, and N2-flushing 2 min before and 2 min after introducing the sample of
tissue, were almost identically effective. The addition of NADH and glucose as external sources of reducing
power was studied and the problems associated with NADH interference in NO2 determination are
discussed. The addition of DCMU did not overcome the need for darkness as it does in higher plants, even
when the photosynthetic O2-production was completely abolished at the same DCMU concentration.
Key words: In situ assay; Nitrate reductase; Ulva rigida
INTRODUCTION
The determination of nitrate reductase (NR) activity in situ has become an alternative
method when the extraction procedure of NR is difficult, or even impossible, without
an inactivation of the enzyme. Metallic ions, phenolic compounds and proteases have
been reported as responsible for NR inactivation in the extraction procedure in different
vegetal tissues (Beevers & Hageman, 1980; Hageman & Reed, 1980). So far attempts
to measure NR activity in vitro in Ulva rigida C. Agardh have failed and the use of
protective compounds such as polyvinylpyrrolidone and cysteine were not effective.
The in situ method has been widely used in higher plants (Jaworski, 1971; Brunetti
& Hageman, 1976) as an alternative method to the in vitro assay, but also as a very
interesting experimental approach to studying some features of the N O f reducing
system in a situation closer to that existing in vivo. The effect on N O r reduction of
different compounds: NADH, FMNH2 (Rhodes & Stewart, 1974; Mauriflo et al.,
Correspondence address: A. Corzo, Department of Ecology, University of Mfilaga, Campus de Teatinos,
29()71 Mfilaga, Spain.
0022-0981/91/$03.50 © 1991 Elsevier Science Publishers B.V. (Biomedical Division)
182
A. CORZO AND F.X. NIELL
1985), various intermediary metabolites of the glycolytic, pentose phosphate, and citric
acid pathways (Klepper et al., 1971) have all been studied using the in situ method.
Canvin & Woo (1979) demonstrated the existence of competition between NOareduction and 0 2 reduction at the end of the respiratory chain by NADH in the dark
using the in situ assay.
The in situ NR assay has reached a high degree of sophistication in higher plants and
is very well understood in general terms. Although it has been performed previously with
seaweed (Dipierro et al., 1977; Weidner & Kiefer, 1981; Davinson & Stewart, 1984;
Brinkhuis et al., 1989), a number of necessary conditions were not fulfilled. It was
performed at low light intensity (Brinkhuis et al., 1989) when the requirement for
darkness was clearly demonstrated in all the cases studied. No care was taken to assure
anaerobic conditions in most of the previous studies despite the fact that it has been
shown to be an important requirement in obtaining the highest possible NR activity
(Canvin & Woo, 1979).
In this paper we describe some of the key conditions for optimising the method in
U. rigida. The effects of light pretreatment are discussed in relation to the use of in situ
NR assay for measuring the N O r metabolic pool as well as in relation to the NR activity
itself. Because it is easy to do, the NR in situ assay should be more effective when
applied to field studies than the in vitro assay which is time-consuming, difficult and
requires more complex equipment and training.
MATERIAL AND METHODS
Ulva rigida was collected in the south of Spain (Algeciras, Cadiz) from a rocky shore.
It was kept in filtered (Whatman GF/C) sea water, which was collected in the same
place, for between 1 and 5 days in a cold chamber (15 ° C) under continuous white light
(120 #mol. m - 2. s - ~). Algae kept in the same conditions for up to 10 days have not
shown any apparent damage.
Before the experiment, samples of 0.16 g of fresh tissue were cut into smaller pieces
and introduced into test-tubes containing 5 ml of the assay medium. Cutting the blade
into smaller pieces increases the reproducibility of the result considerably because it
increased the homogeneity of the assay medium during incubation. After the introduction of the samples, the test-tubes were sealed with rubber stoppers (Aldrich) and
a vacuum (7.9 x 104 Pa) created for 10 min through a needle connected to a pump. To
avoid 02 in the assay medium a second method was used. The medium was flushed
with N 2 for 2 min, and the sample introduced immediately afterwards. The sample plus
medium were bubbled again for 2 min. Finally the test-tubes were quickly sealed. After
vacuum or N 2 treatments, the test-tubes were covered with aluminum foil and incubated
in the dark for 1 h at 30 ° C in a water bath. After that time a sample of 1 ml was removed
from the assay medium and NO2 was determined colorimetrically (Snell & Snell,
1949). All the assays were carried out on duplicate samples of tissue. The composition
NITRATE REDUCTASE IN ULVA RIGIDA
183
of the assay medium was modified in the course of the work when the optimal concentration of each one of its components was found. The final assay medium used was
composed of 30 mM KNO 3, 0.01 mM glucose, 0.1 ~ 1-propanol, 0.05 mM Na-EDTA,
0.1 M phosphate buffer, pH 8.
02 production was measured with a Clark electrode (Rank) (Hansatech DW-2).
Photon Fluence Rate (PFR) was measured with a Quantum Radiometer Li-Cor
(Li-1000 Data Logger) with a spherical sensor (Li-Cor 193 SB). White light was
provided by a Sylvania F 18 W/GRO.
RESULTS AND DISCUSSION
The measurement of NR in situ requires a buffer, a compound able to permeabilise
the membrane, a source of NO3 and a source of reducing power. The assay must be
performed in the dark to avoid the reduction of NO2 mediated by nitrite reductase
which takes electrons directly from reduced ferredoxin. It has also to be done in
anaerobic conditions or at least with low 02 concentration in the medium. This fact has
been attributed to competition for the NADH between nitrate reduction and 02 as the
final acceptor in the respiratory chain (Canvin & Woo, 1979; Beevers & Hageman,
1980; Reed & Canvin, 1982).
In U. rigida the first measurements were performed following the method of Maurifio
et al. (1986), developed for higher plants. The assay medium consisted of 50 mM
KNOa, 1~o 1-propanol, 0.5 mM Na-EDTA, 0.1 M potassium phosphate buffer,
pH 7.5, and the vacuum-infiltration procedure already described was initially used.
PERMEABILISERS AND PHOSPHATE BUFFER
Permeabilisers classically used in the NR in situ assay belong to two categories:
alcohols and surfactants. Although their modes of action have yet to be clarified, the
general opinion is that they operate by increasing membrane permeability therefore
favouring both the entry of NO3 to the reduction site and the exit of NO 2 into the
medium (Mann et al., 1979; Lawrence & Herrick, 1982). Both types of compound have
been used in different plant material with different levels of success. Within alcohols
1-propanol is the most effective in a number of species (Jaworski, 1971; Sym, 1984).
Of the three concentrations of 1-propanol (0.1, 0.5, 1~o) tested in U. rigida, 0.1 ~ was
the best because it produced 40% more activity.
Surfactants such as Tergitol NP-10, Triton X-100 and Neutronix 600 have produced
better results than 1-propanol in some species (Brunetti & Hageman, 1976; Davis &
Ross, 1985). In U. n'gida, however, neither thc substitution of 0.1 ~o l-propanol by
Tergitol NP-10 (0.001, 0.005, 0.1 ~o) nor by Triton X-100 (0.001, 0.005, 0.01 ~ ) in the
assay medium increased the rate of NR activity (results not shown). Toluene dissolved
in ethanol has also been used in the in situ NR assay as a permeabiliser (Choudary,
1984) giving better results than n-propanol in some species of phytoplankton (Hochman
184
A. CORZO AND F.X. NIELL
et al., 1986) but worse results in Monoraphidium braunii and Peridinium cinctum (R. Plasa
& W.R. Ullrich, pers. comm.).
Two dilutions (1/2 and 1/10) of the initial buffer (0.1 M potassium phosphate
buffer + 0.5 mM Na-EDTA) were tested without obtaining better results. Thenceforth
0.1 M phosphate buffer + 0.5 mM Na-EDTA and 0.1 ~o 1-propanol were used.
KNO 3 OPTIMUM
The addition of NO3 to the assay medium enhanced in situ activity considerably and
additionally standardised the N O r production rate which would otherwise be very
much dependent on NO3 intracellular concentration and therefore on the pre-experimental treatment (Fig. 5). In Ulva rigida the NR activity is already saturated at 30 mM
KNO3 (Fig. 1). A NO~- concentration higher than optimal produced an inhibitory effect
on NR activity. Simiiar responses to increasing NO3 concentration have been found
in higher plants (Timpo & Neyra, 1983; Davis & Ross, 1985; Mauriflo et al., 1986) and
in the unicellular green alga Monoraphidium bmunii (Corzo, in prep.). However, no
satisfactory explanation has been suggested for this fact. All remaining experiments were
done with 30 mM KNO 3 in the assay medium.
0.3
7
:~
0.2
7
I~¢NI
z
o
[]
/"
0.1
E
1"
t
I
0
z
I
I0
.
.
I
I
I
[
20
30
40
50
I
60
KNO~ (mM)
Fig. I. NR activity (#mol NO~- .g- '. (w.w.) h- ~) dependence on KNO3 concentration in the incubation
medium. Means of two replicates. SD represented as a bar when greater than the symbol size.
pH
The external pH optimum for NR in situ assay in U. rigidawas found at pH 8 (Fig. 2).
As far as we know this is the highest pH optimum ever reported for the in situ method
and it is similar to the pH optimum for in vitro NR activity from Porphyra yezoensis
(Shigeru et al., 1979). There is a clear difference with respect to the optimum pH found
NITRATE REDUCTASE IN UL VA RIGIDA
185
0.3
7
7~
'[/
'Q"
z- -
0.I
[]
0
.../.,~._.1 /
6
,
n
,
7
n
8
,
9
pH
Fig. 2. Effect of pH on the NR activity (#mol NOr "g-~'(w.w.) h-~). Means of four replicates. SD
represented as a bar when greater than the symbol size.
in higher plants between 7 and 7.5 (Sym, 1984; Maurifio et al., 1986) and that found
in U. rigida. It is interesting to point out the proximity of that value to the pH of sea
water (8.2).
NADH AND GLUCOSE
NADH as been widely used as an external source of reducing power in the determination of NR activity in vitro and in situ. But the use of NADH presents a technical
problem because of its interference with the synthesis of the azo dye in the NO2
determination when sulphanilamide and N-naphtylethylenediamine are used (Medina
& Nicholas, 1957; Zumft et al., 1983; Davies & Ross, 1985). Different methods have
been used to avoid this inhibition (Medina & Nicholas, 1957; Hochman et al., 1986).
However, we have not had much success with either method. When the N O r concentration in the assay medium is lower than 30 #M, neither the method of Medina &
Nicholas (1957) nor Hochman's method avoided the NADH inhibition of the chromatic
reaction. Scholl et al. (1974) reported a different method which consists of the use of
phenazine methosulfate to oxidized NADH. Although it has been used mainly in the
in vitro NR assay (Scholl et al., 1974; Scheideler & Ninnemann, 1986), it could be an
interesting approach to in situ NP. assay.
The NAD(P)H is the physiological source of reducing power for N O r reduction.
In vivo, NAD(P)H may have two different origins, it may be produced by photosynthesis, or by respiration. When NR activity is determined by the in situ assay (incubation
in the dark), the enzyme has only two possible sources of reducing power: NADH
produced by respiratory metabolism using reserves already existing in the cell and/or
some metabolite externally supplied. Besides NADH and F M N H : (Rhodes & Stewart,
186
A. CORZO AND F.X. NIELL
1974; Mauriflo et al., 1985), different substrate and intermediary ofglycolysis as glucose
6-P, fructose diphosphate, 1-3-diphosphoglycerate have been used to guarantee a source
of reducing power (Klepper et al., 1971).
In U. n'gida, the addition of glucose to in situ NR assay medium as a source of NADH
in the dark, increases the NR activity by 69~ with respect to a control without any
external source of reducing power, and by 17Yo with respect to a control with 0.12 mM
NADH (optimal concentration when Medina & Nicholas's method was used). The
differences, using three glucose concentrations ( 10 - 3, 10 - 2, 10 - ~mM), were, however,
not statistically significant. Thenceforth 10-2 mM glucose was used as routine in the
assay medium in order to rule out a limitation in NO3 reduction rate as a consequence
of a lack of reducing power.
VACUUM AND N2-FLUSHING
The in situ assay of NR activity requires anaerobic conditions to obtain the highest
possible rate of nitrate reduction. The lower level of NR measured in the presence of
02 has been explained as a competition between nitrate reduction and 0 2 reduction into
H20 at the end of the respiratory chain, by the reducing power available in the cytoplasm
(Canvin & Woo, 1979; Beevers & Hageman, 1980; Reed & Canvin, 1982). Both vacuum
infiltration for 10 min and N 2 flushing were almost equally effective (6.8 and 7 times
higher respectively than the control), and in addition the use ofboth methods at the same
time did not significantly increase the rate of NR activity. From a practical point of view,
however, we decided to employ the second method (N 2 flushing) because it is less
time-consuming.
DCMU
In higher plants, when DCMU is used in in situ assay, the requirement for darkness
to observe an NO~- accumulation can be obviated (Miflin, 1974; Neyra & Hageman,
1974, Atkins & Canvin, 1975; Klepper, 1979; Maurifio et al., 1986) because the electronic flux up to ferredoxin, which is supposed to be the direct electronic donor of the
nitrite reductase, is blocked by DCMU. In U. rigida, however, the use of DCMU at the
same concentration, or even higher than that at which the photosynthetic production
of 02 was already inhibited (Fig. 3), was almos: completely ineffective in avoiding an
NO~- reduction (Table I) in spite of the fact that NO~- reduction and NO~- reduction
in green algae and in higher plants are considered as very similar.
Apparently NR is not directly affected by DCMU because when the NR activity was
assayed in the presence of DCMU, in the dark, rates were higher than in the control
if glucose was provided or lower (70% of control) if NADH was provided externally.
In both cases, however, it was higher than that obtained in the presence of DCMU when
the assay was performed in the light (150 #tool. m - 2. s - ~). The measurement of NO3
consumption and NH~ production in the in situ NR assay in the presence of DCMU
both in the light and in darkness did not provide any clear additional information. We
N I T R A T E R E D U C T A S E IN UL VA RIGIDA
187
-S
5xIO
M
DCMU
LighI off
O
E
If
Imin
Fig. 3. D C M U (5 x 10- 5 M) inhibition of 02 production in U. rigida. The experiment was performed at
PFR of 730/~mol • m - 2. s - ~
TABLE I
Effects of 5 x 10-5 M D C M U on N R activity expressed as 9o of control (incubation in dark without
D C M U ) in dark and in light (150 #mol. m -2" S - i ) using exogenous N A D H (0.12 m M ) or glucose (0.01
raM) as an external source of reducing power (means of four replicates _+so).
÷ Glucose
+ NADH
Dark
Dark ÷ D C M U
Light + D C M U
100
100
122 _ 2
70 +_ 2
21 _ 16
19 _+ l0
have no explanation for such a difference in response to DCMU between higher plants
and U. rigida and further investigation is necessary.
T I M E C O U R S E O F NO 2
PRODUCTION
Figure 4 illustrates the time course of NO2- production in the in situ NR assay
performed either without exogenous NO3 or with 30 mM KNO3. The plateau level was
almost reached in 30 min and after 2 h the level of NO2 decreased. This response was
shown in both cases; both when 30 mM NO~ was added and when the only source
of NO~ was the internal pool. Such decay suggests the existence of a certain rate of
NO2 reduction even in the dark, which may cause an underestimation of the existing
NR activity. The use of a shorter incubation time, however, should overcome this
trouble.
The plateau level (cessation of NO2 accumulation) when 30 mM N O r was added
to the assay medium was twice as high as without an external supply of NO 3 (Fig. 4),
but this difference is dependent upon the internal NO 3 pool size and therefore upon
pre-experimental treatment. The plateau level determined by in situ NR assay when no
NO3 is supplied has been proposed as a method for determining the size of the
metabolic NO3 pool in higher plants (Ferrari et al., 1973; SteingrOver et al., 1986).
188
A. CORZO AND F.X. NIELL
0.30
0.24
.'-
+NO~ . 0 - -
o.18
'o"
z
0.12
=
0.06
I
+NO~
/o.o--'-o\
No;
]
0
I
i
=./e-=
|
I
l
60
120
180
•
I
240
•
I
.
300
t (min)
Fig. 4. Time course of NO~- production in the in situ assay. Assay medium containing 30 mM KNO 3 ([]).
Assay medium without exogenous NO~- ( i ) ; the arrow indicates the time at which 30 mM KNO3 was
added to the incubation medium. SD represented as a bar when greater than the symbol size.
Despite that, it has been shown that this level changes depending on the availability of
NO£ for the plants before the assay. It cannot be unambiguously established whether
the NR activity measured through the in situ assay is only being limited by the cytoplasmic NO3 concentration. Since the estimation of the metabolic NO£ pool requires the
determination of the plateau level instead of the initial rate, and a longer incubation time
is therefore needed, the lack of reducing power or a further inactivation of NR by the
assay conditions themselves could cause an underestimation of the metabolic NO~pool. The decay in the plateau level after 2 h (Fig. 4) could be a consequence of these
kinds of limiting processes previously mentioned. The addition after 3 h of 30 mM
KN03 to the assay, which originally received no exogenous N O £ , did not re-establish
NO2 production. Thus a limitation of NR activity as a consequence of lack of substrate
was ruled out and, therefore, the use of the plateau level as a good estimate of endogenous NO3 pool at that stage.
NR activity measurable by in situ assay was dependent on the light condition in which
the alga was kept prior to the assay. The results in Fig. 5 show the effect on the
NR activity of keeping the alga in the dark for 15 h prior to the experiment or in white
light (300 #mol. m - 2. s - ~) in sea water with low NO3 concentration (0.8 #M). When
U. rigida was kept in the dark before the NR assay, some NR activity could be detected
even in the absence of an external NO3 supply in the assay medium (the source of N 0 3
in this case was the internal pool). On the other hand, when the alga was kept in the
light before the experiment no NO2 production could be detected in NO3 free assay
medium for 90 min. The simplest explanation is that in this case the N 0 3 internal pool
was exhausted as a consequence of the higher nitrogen demand in the light. In U. rigida
NITRATE REDUCTASE IN UL VA RIGIDA
189
0.4
"~
0.3
";'~
,~,
0.2
/(:!
+NO~
0
[]
T
°
~m
o
E
0.1
~"
,.....m-~m/_m~,i,/°
~i~_o--n--~-.
0
30
0
60
90
.
,
120
.
,
150
.
,
.
180
t (min)
Fig. 5. Time course of NO2 production in the in situ NR assay without exogenous NO3 addition. 15 h
before the experiment U. rigida was kept in natural sea water with low NO; concentration (0.8 gM) in white
light (300 #mol .m -2. s-~) (I-1); or in the dark (m). After 90 min 30 mM KNO3 was added to the
incubation medium in both cases. SD deviation represented as a bar when greater than the symbol size.
the close dependence of the NO3- internal pool on light intensity has been shown
elsewhere (Corzo & Niell, in press).
The addition of NO3 after 90 min showed other important features of the NO3reduction system in U. rigida. In both cases there was an immediate and clear increase
of the NO£ production, which was higher when the alga was kept in the light before
the experiment, and reached a higher plateau level (Fig. 5). It has been shown that
NR activity increases in the light both in higher plants and in unicellular algae (Hattori,
1962; Beevers & Hageman, 1972; Aparicio et al., 1976; Azuara & Apaficio, 1983)
in vitro and in situ. Similarly, the NR of Ulva rigida is also activated by light (Corzo,
in prep.). A higher level of NR activity as a consequence of previous light pretreatment
could, therefore, account for both the higher initial rate of NO2 production and the
higher plateau level. Additionally, a higher NAD(P)H/NAD(P) ratio as a consequence
of previous light pretreatment could account for part of the observed response.
ACKNOWLEDGEMENTS
This work was supported by a grant (PB86-0677) from the Comisi6n Asesora de
Investigaci6n Cientifica y T6cnica (Spain).
REFERENCES
Aparicio, P.J., J. M. Roldan & F. Calero, 1976. Blue light photoreactivatiol of nitrate reductase from green
algae and higher plants. Biochem. Biophys. Res. Commun., Vol. 70, pp. 1071-1077.
190
A. CORZO AND F.X. NIELL
Atkins, C. A. & D.T. Canvin, 1975. Nitrate, nitrite and ammonia assimilation by leaves: effects ofinhibitors.
Planta, Vol. 123, pp. 41-51.
Azuara, M.P. & P.J. Aparicio, 1983. In vivo blue-light activation of Chlamydomonas reinhardii nitrate
reductase. Plant Physiol., Vol. 71, pp. 286-290.
Beevers, L. & R.H. Hageman, 1972. The role of light in nitrate metabolism in higher plants. In: Photophysiology, edited by A.C. Giese, Academic Press, New York, Vol, 7, pp. 85-113.
Beevers, L. & R.H. Hageman, 1980. Nitrate and nitrite reduction. In: The biochemistry of plants, edited by
B.J. Miflin, Academic Press, New York, Vol. 5, pp. 115-168.
Brinkhuis, B. H., L. Renzhi, W. Chaoyuan & J. Xun-Sen, 1989. Nirite uptake transients and consequences
for in vivo algal nitrate reductase assays. J. Phycoi., Vol. 25, pp. 539-545.
Brunetti, N. & R. H. Hageman, 1976. Comparison of in vivo and in vitro assays ofnitrate reductase in wheat
(Triticum aestivum L. ) seedlings. Plant Physiol., Vol. 58, pp. 583-587.
Canvin, D.T. & K. C. Woo, 1979. The regulation of nitrate reduction in spinach leaves. Can. J. Bot., Vol. 57,
pp. 1155-1160.
Choudary, P.V., 1984. A simple micromethod for rapid kinetic analyses of yeast enzymes in situ. Anal.
Biochem., Vol. 138, pp. 425-429.
Corzo, A. A. & F.X. Niell. Inorganic nitrogen metabolism in Ulva rigida illuminated with blue light. Mar.
BioL, in press.
Davies, H.V. & H.A. Ross, 1985. Factors affecting in vivo and in vitro determinations of nitrate reductase
activity in potato leaves. J. Plant Physiol., Vol. 119, pp. 1-7.
Davinson, I. R. & W. D. P. Stewart, 1984. Studies on nitrate reductase activity in Laminaria digitata (Huds.)
Lamour. I. Longitudinal and transverse profiles of nitrate reductase activity within the thallus. J. Exp.
Mar. Biol. Ecol., Vol. 74, pp. 201-210.
Dipierro, S., C. Perrone & G.P. Felicini, 1977. In vivo nitrate reductase assay in Petroglossum nicaeense
(Duby) Schotter (Rhodophyta, Phyllophoraceae). Phycologia, Vol. 16, pp. 179-182.
Ferrari, T. E., O. C. Yoder & P. Filner, 1973. Anaerobic nitrite production by plant cells and tissues: evidence
for two nitrate pools. Plant PhysioL, Vol. 51, pp. 423-431.
Hageman, R.H. & A.J. Reed, 1980. Nitrate reductase from higher plants. Methods EnzymoL, Vol. 69,
pp. 270-280.
Hattori, A., 1962. Light-induced reduction of nitrate, nitrite and hydroxylamine in a blue-green alga,
Anabaena cylindrica. Plant Cell. Physiol., Vol. 8, pp. 327-337.
Hochman, A., A. Nissany, D. Wynne, B. Kaplan & T. Berman, 1986. Nitrate reductase: an improved assay
method for phytoplankton. J. Plankt. Res., Vol. 8, pp. 385-392
Jaworski, E.G., 1971. Nitrate reductase assay in intact plant tissues. Biochem. Biophys. Res. Commun.,
Vol. 43, pp. 1274-1279.
Klepper, L., D. Flesher & R.H. Hageman, 1971. Generation of reduced nicotinamide adenine dinucleotide
or nitrate reduction in green leaves. Plant Physiol., Vol. 48, pp. 580-590.
Klepper, L.A., 1979. Effects of certain herbicides and their combinations on nitrate and nitrite reduction.
Plant Physiol., Vol. 64, pp. 273-275.
Lawrence, J. M. & H.E. Herrick, 1982. Media for in vivo nitrate reductase assay of plant tissues. Plant Sci.
Leit., Vol. 24, pp. 17-26.
Mann, A. F., D. P. Hucklesby & E.J. Hewitt, 1979. Effect of aerobic and anaerobic conditions on the in vivo
nitrate reductase assay in spinach leaves. Pianta, Vol. 146, pp. 83-89.
Maurifio, S.G., C. Echevarria, M.A. Vargas & J.M. Maldonado, 1985. Effect of freezing and thawing on
in situ nitrate reductase activity in spinach and maize leaves. J. Plant PhysioL, Vol. 120, pp. 409-417.
Maurifio, S. G., C. Echevarria, J. A. Mejias, M. A. Vargas & J. M. Maldonado, 1986. Properties ofthe in vivo
nitrate reductase assay in maize, soybean, and spinach leaves. J. Plant Physiol., Vol. 124, pp. 123-130.
Medina, A. & D.J.D. Nicholas, 1957. Interference by reduced pyridin¢ nucleotides in the diazotation of
nitrite. Biochim. Biophys. Acta, Vol. 23, pp. 440-442.
Miflin, B.J., 1974. Nitrite reduction in leaves; studies on isolated chloroplasts. Planta, Voi. 116, pp. 187-196.
Neyra, C. A. & R. H. Hageman, 1974. Dependence of nitrite reduction on electron transport in chloroplasts.
Plant Physiol., Vol. 54, pp. 480-483.
Reed, A.J. & D.T. Canvin, 1982. Light and dark controls of nitrate reduction in wheat (Triticum aestivum
L. ) protoplasts. Plant Physiol., Voi. 69, pp. 508-513.
Rhodes, D. & G.R. Stewart, 1974. A procedure for the in vivo determination of enzyme activity in higher
plant tissue. Planta, Vol. 118, pp. 133-144.
NITRATE REDUCTASE IN ULVA RIGIDA
191
Scheideler, L. & H. Ninnemann, 1986. Nitrate reductase activity test: phenazine methosulfate-ferricyanide
stop reagent rep!aces postassay treatment. Anal Biochem., Vol. 154, pp. 29-33.
Scholl, R. L., J. E. Harper & R.H. Hageman, 1974 Improvements of nitrite color development in assays of
nitrate reductase by phenazine methosulfate and zinc acetate. Plant Physiol., Vol. 53, pp. 825-828.
Shigeru, A., T. Ikawa, T. Oohusa & K. Nisizawa, 1979. Some enzymic properties of nitrate reductase from
Porphyra yezoensis f. narawaensis. Bull. Jpn. Soc. $ci. Fish., Vol. 45, pp. 919-924.
Snell, F. D. & C.T. Snell, 1949. Colorimetric Methods of Analysis. 3rd edition, Van Nostrand, Princeton, New
Jersey, Vol. 2, 804 pp.
Steingr6ver, E., P. Ratering & J.Siesling, 1986. Daily changes in uptake, reduction and storage of nitrate
in spinach grown at low light intensity. Physiol. Plant., Vol. 66, pp. 550-556.
Sym, G.J., 1984. Optimisation of the in vivo assay conditions for nitrate reductase in Barley (Hordeum
vulgare L. cv. Igri). J. Sci. Food Agric., Vol. 35, pp. 725-730.
Tim~o, E. E. & C.A. Neyra, 1983. Expression ofnitrate and nitrite reductase activities under various forms
of nitrogen nutrition in Phaseolus vulgaris L.. Plant Physiol., Vol. 72, pp. 71-75.
Weidner, M. & H. K'~efer, 1981. Nitrate reduction in the marine brown alga Giffordia mitchellae (Harv.) Ham.
Z. PflanzenphysioT., Vol. 104, pp. 341-351.
Zumft, W.G., F. Castillo & K.M. Hartman, 1983. Flavin-mediated photoreduction of nitrate by nitrate
reductase of higher plants and microorganisms. In, The blue light syndrome, edited by H. Senger, SpringerVerlag, Berlin.