6---Metabolism_1989_The-Chlamydomonas-Sourcebook

Prévia do material em texto

6 Metabolism 
Introduction 
Despite the ease with which Chlamydomonas can be cultured and ma-
nipulated, detailed biochemical and physiological studies have been 
done with relatively few enzyme systems in this organism, and molecu-
lar analysis of genes coding for specific enzymes is just beginning, in 
contrast to some other areas of Chlamydomonas research such as flagel-
lar biogenesis, chloroplast structure and function, and the mating reac-
tions, in which biochemical, genetic, and molecular studies can be inte-
grated into a coherent view. The present chapter therefore deals with 
several diverse topics, beginning with a summary of investigations of 
specific enzymes (Table 6.1). The best-characterized metabolic path-
ways, including carbon metabolism, respiration and chlororespiration, 
hydrogenase, lipid biosynthesis, nitrogen assimilation, and arginine bio-
synthesis, are then discussed in some detail. The chapter concludes with 
tabular summaries of the metabolic inhibitors and herbicides which have 
been tested on Chlamydomonas. Algal Physiology and Biochemistry, 
edited by Stewart (1974), is still a good summary of basic metabolic 
processes in the algae as a group, and relevant background material will 
be found there. An older book, Physiology and Biochemistry of Algae, 
edited by Lewin (1962), provides a summary of the early literature. 
Acetate Flagellates 
Pringsheim (1937) proposed the term "acetate flagellates" to describe 
the colorless Polytoma, which grows well on acetate as sole carbon 
source but cannot use glucose. Later authors (Hutner and Provasoli, 
1951 ; L loyd and Cantor, 1979) have extended the designation to include 
an assortment of both green and colorless cells, some of which can also 
use pyruvate or lactate. The acetate flagellates generally have plasma 
membranes with low permeability to most organic substrates, with only 
small, lipid-soluble molecules showing good penetration. A s a group, the 
acetate flagellates are able to tolerate low 0 2 tension and high levels of 
C 0 2 . Pringsheim (1946b) used pieces of cheese covered with soil and 
water to enrich cultures in colorless acetate flagellates, which grew well 
in the relatively high levels of fatty acids and alcohols produced by 
bacteria in this milieu. In nature, acetate flagellates are found in similar 
217 
218 6. Metabolism 
EC number Enzyme Reference 
EC 1.1.1.1 Alcohol dehydrogenase Grondai et al. (1983); Kreuzberg et al. (1987) 
EC 1.1.1.3 Homoserine dehydrogenase Vincze and Dénes (1968, 1973) 
EC 1.1.1.25 Shikimic acid dehydrogenase Berlyn et al. (1970) 
EC 1.1.1.26 Glyoxylate reductase Hess and Tolbert (1967a); Zelitch and Day 
(1968); Bruin et al. (1970); Husic and 
Tolbert (1987a) 
EC 1.1.1.26? "Glycolate oxidase" Zelitch and Day (1968) 
EC 1.1.1.27 Lactate dehydrogenase Kreuzberg et al. (1987); Husic and Tolbert 
(1987b) 
EC 1.1.1.28 D-lactate dehydrogenase D. W. Husic and Tolbert (1985) 
EC 1.1.1.29 Hydroxypyruvate reductase (glycerate dehy- Stabenau (1974); Husic and Tolbert (1987a) 
drogenase) 
EC 1.1.1.37 Malate dehydrogenase Thomas and Delcarpio (1971); Frankel and 
Jones (1980) 
EC 1.1.1.42 Isocitrate dehydrogenase Hess and Tolbert (1967a); Ramaley and 
Hudock (1973); Foo and Badour (1977) 
EC 1.1.1.43 Phosphogluconate dehydrogenase Herbert et al. (1979); Hipkin and Cannons 
(1985); Klein (1986) 
EC 1.1.1.49 Glucose-6-phosphate dehydrogenase Herbert et al. (1979); Hipkin and Cannons 
(1985); Klein (1986) 
EC 1.1.99.14 Glycolate dehydrogenase Nelson and Tolbert (1969, 1970); Bruin et al. 
(1970); Cooksey (1971); Paul and Volcani 
(1976); Husic and Tolbert (1987b) 
EC 1.2.1.10 Aldehyde dehydrogenase Kreuzberg et al. (1987) 
EC 1.2.1.- Glyceraldehyde-3-phosphate dehydrogenase Hudock and Bart (1969) 
EC 1.2.1.12 Glyceraldehyde-3-phosphate dehydrogenase Klein (1986) 
( N A D ) 
EC 1.2.1.13 Glyceraldehyde-3-phosphate dehydrogenase Klein (1986) 
(NADP) 
EC 1.2.1.37 Xanthine dehydrogenase Fernandez and Cârdenas (1981a); Pérez-
Vicente et al. (1988) 
EC 1.2.1.38 Acetylglutamyl-phosphate reductase Strijkert and Sussenbach (1969) 
EC 1.2.4.1 Pyruvate dehydrogenase Kreuzberg et al. (1987) 
EC 1.4.1.1 Alanine dehydrogenase Kates and Jones (1964c, 1967); Frankel and 
Jones (1980) 
EC 1.4.1.3 Glutamate dehydrogenase Kates and Jones (1964c, 1967); Hudock and 
Bart (1967); Kivic et al. (1969); Thomas 
and Delcarpio (1971); Frankel and Jones 
(1980); Paul and Cooksey (1981a); Culli-
more and Sims (1981a) 
EC 1.4.1.14 Glutamate synthase ( N A D H ) Cullimore and Sims (1981a,b): Marquez et al. 
(1984, 1986a) 
EC 1.4.7.1 Glutamate synthase (ferredoxin) Cullimore and Sims (1981a,b); Marquez et al. 
(1984, 1986a,b); Galvân et al. (1984) 
EC 1.6.6.2 Nitrate reductase Barea and Cârdenas (1975); Franco et al. 
(1984a); also see text 
EC 1.7.3.3 Urate oxidase Pineda et al. (1984a,b) 
EC 1.7.7.1 Nitrite reductase Barea and Cârdenas (1975); Florencio and 
Vega(1982, 1983a) 
T a b l e 6.1 E n z y m e s R e p o r t e d or Charac te r i zed f rom Chlamydomonas
8 
Acetate Flagellates 
T a b l e 6.1 {continued) 
219 
EC number Enzyme Reference 
EC 1.9.3.1 Cytochrome c oxidase Klein (1986); Husic and Tolbert (1987b) 
EC 1.11.1.6 Catalase Bruin et al. (1970); Chua (1971); Stabenau 
(1974); Paul and Volcani (1976); Klein 
(1986) 
EC 1.11.1.- Ascorbate peroxidase Kow et al. (1982) 
EC 1.14.11.2 Prolyl hydroxylase Blankenstein et al. (1986) 
EC 1.18.3.1 Hydrogenase Roessler and Lien (1984a-c); see text 
EC 2.1.1.45 Thymidylate synthase Vandiver and Fites (1979) 
EC 2.1.2.1 Serine hydroxymethyl transferase Bruin et al. (1970) 
EC 2.1.3.2 Aspartate carbamoyltransferase Kates and Jones (1967) 
EC 2.1.3.3 Ornithine carbamoyltransferase Kates and Jones (1967); Strijkert and Sussen-
bach (1969); Holden and Morris (1970) 
EC 2.2.1.1 Transketolase Salvucci and Ogren (1985) 
EC 2.3.1.8 Phosphotransacetylase Kreuzberg et al. (1987) 
EC 2.3.1.15 Glycerophosphate acyltransferase Jelsema et al. (1982); Michaels et al. (1983) 
EC 2.3.1.35 Glutamate acetyltransferase Staub and Dénes (1966); Dénes (1971c) 
EC 2.3.1.42 Dihydroxyacetonephosphate acyltransferase Jelsema et al. (1982) 
EC 2.3.1.54 Pyruvate formate-lyase Kreuzberget al. (1987) 
EC 2.3.1.- Lysophosphatidate acyltransferase Jelsema et al. (1982); Michaels et al. (1983) 
EC 2.4.1.- Galactosyl transferase Mendiola-Morgenthaler et al. (1985b) 
EC 2.4.1.1 Phosphorylase Klein (1986) 
EC 2.5.1.19 3-Enolpyruvylshikimate-5-phosphate syn- Berlyn et al. (1970) 
thase 
EC 2.6.1.4 Glycine aminotransferase Bruin et al. (1970) 
EC 2.6.1.11 Acetylornithine aminotransferase Siidi and Dénes (1967a); Strijkert and Sussen-
bach (1969); Dénes (1971d) 
EC 2.6.1.13 Ornithine aminotransferase Südi and Dénes (1967b) 
EC 2.7.1.1 Hexokinase Delvalle and Asensio (1978) 
EC 2.7.1.2 Glucokinase Uryson and Kulaev (1970) 
EC 2.7.1.11 Phosphofructokinase Klein (1986) 
EC 2.7.1.19 Phosphoribulokinase Moll and Le vine (1970); Salvucci and Ogren 
(1985) 
EC 2.7.1.21 Thymidine kinase Swinton and Chiang (1979) 
EC 2.7.1.25 Adenylylsulfate kinase Schwenn and Jender (1981); Jender and 
Schwenn (1984); Schwenn and Schriek 
(1984) 
EC 2.7.1.31 Glycerate kinase Bruin et al. (1970) 
EC 2.7.1.40 Pyruvate kinase Klein (1986) 
EC 2.7.1.71 Shikimate kinase Berlyn et al. (1970) 
EC 2.7.2.1 Acetate kinase Kreuzberg et al. (1987) 
EC 2.7.2.8 Acetylglutamate kinase Farago and Dénes (1967, 1969a,b); Dénes 
(1971b) 
EC 2.7.5.1 Phosphoglucomutase Herbert et al. (1979); Klein (1986) 
EC 3.1.1.- α-Esterase Thomas and Delcarpio (1971) 
EC 3.1.3.1 Alkaline phosphatase Lien and Knutsen (1972, 1973a); Loppes 
(1978); Matagne et al. (1976a,b); Patni et al. 
(1977) 
EC 3.1.3.2 Acid phosphatase Matagne et al. (1976a,b); Patni et al. (1977) 
EC 3.1.3.11 Fructose biphosphatase Klein (1986) 
(continued) 
220 6. Metabolism 
EC number Enzyme Reference 
EC 3.1. 3.18 Phosphoglycolate phosphatase Hess and Tolbert (1967a); Nelson and Tolbert 
(1969); Bruin et al. (1970); H.D. Husic and 
Tolbert (1984, 1985) 
EC 3.1. .4.17 Cyclic nucleotide phosphodiesterase Amrhein and Filner (1973); Fischer and 
Amrhein (1974) 
EC 3.1. .6.1 Arylsulfatase Lien and Schreiner (1975); de Hostos et al 
(1988) 
EC 3.2. .1.1 α-Amylase Levi and Gibbs (1984) 
EC 3.2. 1.- Amylase Klein (1986) 
EC 3.4. 11.- Aminopeptidases Thomas and Delcarpio (1971); Lang et al. 
(1979) 
EC 3.5. 1.1 Asparaginase Paul and Cooksey (1979, 1981a,b); Paul 
(1982) 
EC 3.5. 1.4 Amidase Gresshoff (198la,b); Hodson and Gresshoff 
(1987) 
EC 3.5. 3.6 Arginine deiminase Sussenbach and Strijkert (1969b, 1970) 
EC 4.1. 1.1 Pyruvate decarboxylase Kreuzberg et al. (1987) 
EC 4.1. 1.31 Phosphoenol pyruvate carboxylase Kates and Jones (1967); Chen and Jones 
(1970, 1971); Scrutton and Fatebene (1975); 
Klein (1986) 
EC 4.1. 1.39 Ribulose bisphosphate carboxylase/oxygenase Givan and Criddle (1972); Iwanij et al. (1974); 
Nelson and Surzycki ( 1976a,b); Spreitzer et 
al. (1982) 
EC 4.1. 2.13 Fructose bisphosphate aldolase Guerrini et al. (1971a); Salvucci and Ogren 
(1985); Klein (1986) 
EC 4.1. 3.1 Isocitrate lyase Foo and Badour (1977); Wiseman et al. 
(1977a) 
EC 4.1. .3.18 Acetolactate synthase Hartnett et al. (1987) 
EC 4.2. .1.1 Carbonate dehydratase (carbonic anhydrase) Bundy and Cote (1980); Toguri et al. (1984, 
1986); Coleman et al. (1984); Yang et al. 
(1985); Bundy (1986) 
EC 4.2. .1.10 Dehydroquinate dehydratase Berlyn et al. (1970) 
EC 4.2, .1.11 Phosphopyruvate hydratase Klein (1986) 
EC 4.2 .99.8 Acetyl serine sulfhydrylase Leon et al. (1987) 
EC 4.3. .2.1 Argininosuccinate lyase Strijkert and Sussenbach (1969); Holden and 
Morris (1970); Matagne and Schlosser 
(1977); Farrell and Overton (1987) 
EC 4.6. .1.1 Adenylate cyclase Hintermann and Parish (1979) 
EC 4.6 .1.3 Dehydroquinate acid synthase Berlyn et al. (1970) 
EC 4.7 .2.1 Acetate kinase (acetate phosphotransferase) Farago and Dénes (1968) 
EC 5.3 .1.9 Glucosephosphate isomerase Patni et al. (1977); Herbert et al. (1979); 
Klein (1986) 
EC 5.4 .99.5 Chorismate mutase Zurawski and Brown (1975) 
EC 6.3 .1.2 Glutamine synthetase Cullimore (1981); Cullimore and Sims (1981a); 
Paul and Cooksey (1981a) 
EC 6.3 .4.5 Argininosuccinate synthetase Strijkert et al. (1973) 
EC 6.3 .4.6 Urea carboxylase (hydrolyzing) Hodson et al. (1975); Semler et al. (1975); 
Maitz et al. (1982); Hodson and Gresshoff 
(1987) 
Nucleases, methylases, and other enzymes of D N A synthesis and repair are listed in Chapter 9. 
T a b l e 6.1 (continued) 
Acetate Flagellates 221 
metabolite-rich environments, including sewage, and species of Chlamy-
domonas are among these. 
Not all species of Chlamydomonas are able to grow in the dark on 
acetate, however (Table 6.2), and undoubtedly one of the principal rea-
sons for the ascendancy of C. reinhardtii over C. eugametos and C. 
moewusii as a research organism is its ability to do so, making possible 
the isolation of numerous nonphotosynthetic mutants. Chlamydomonas 
eugametos and C. moewusii are nevertheless capable of using exoge-
nous acetate in the light (J. C. Lewin, 1950; Bernstein, 1968; Der and 
Gowans, 1972; Gowans, 1976a), and mutants that require acetate or 
T a b l e 6.2 Uti l izat ion of A c e t a t e for Heterot rophic Growth by 
Chlamydomonas S p e c i e s
9 
Species Reference 
Growth in the dark on acetate 
C. angulosa Hellebust et al. (1982) 
C. debaryana Lewin (1954b) 
C. dorsoventralis
h 
Lucksch (1933) 
C. dysosmos Lewin (1954b); Haigh and Beevers (1962) 
C. globosa Chlamydomonas Genetics Center unpubl. 
C. komma Chlamydomonas Genetics Center unpubl. 
C. monoica Lucksch (1933) 
C. pallens Pringsheim (1962) 
C. pseudagloë Lucksch (1933) 
C. pseudococcum Lucksch (1933) 
C. pulchra Lucksch (1933) 
C. Pulsatilla (New Brunswick) Hellebust and Le Gresley (1985) 
C. reinhardtii Sager and Granick (1953) 
C. spreta Droop and McGill (1966); Turner (1979) 
C. subglobosa Lucksch (1933) 
Poor or no growth in dark on acetate 
C. chlamydogama Hutner and Provasoli (1951) 
C. elliptica var. britannica Chlamydomonas Genetics Center unpubl. 
C. eugametos Wetherell (1958) 
C. euryale Coughlan (1977) 
C. frankii Chlamydomonas Genetics Center unpubl. 
C. humicola Lucksch (1933) 
C. incerta Chlamydomonas Genetics Center unpubl. 
C. intermedia Chlamydomonas Genetics Center unpubl. 
C. iyengarii Chlamydomonas Genetics Center unpubl. 
C. melanospora Lewin (1975) 
C. moewusii J. C. Lewin (1950); Bernstein (1968) 
C. orbicularis Chlamydomonas Genetics Center unpubl. 
C. philotes Lewin (1957a) 
C. Pulsatilla (Finland, Scotland) Droop (1961); Turner (1979) 
C. smithii Chlamydomonas Genetics Center unpubl. 
C. typica Chlamydomonas Genetics Center unpubl. 
" For further information, see Droop (1974). 
h
 The stock of C. dorsoventralis (UTEX 228) in the Chlamydomonas Genetics Center collection 
does not grow on acetate. 
222 6. Metabolism 
other organic substrates for growth in the light have been isolated in 
these species (Chapter 11). A few species, such as C. nakajaurensis 
(Bennett and Hobbie, 1972), C. acidophila (Erlbaum, 1968), C. pseudag-
loë, and C. pseudococcum (Lucksch, 1933), have been reported to be 
able to use glucose as their sole carbon source, but C. reinhardtii cannot 
do this. Utilization of exogenous ribose by C. reinhardtii was reported 
by Patni et al. (1977), but studies by Sager and Granick (1953) indicated 
that this compound would not support growth in the dark. The other 
compounds tested by Sager and Granick that were nontoxic in the light 
but did not support growth of C. reinhardtii in the dark were as follows: 
glucose, galactose, sucrose, lactose, maltose, mannose, D -xylose , L -
arabinose, ethanol, glycerol, mannitol, formate, glycerophosphate, pro-
pionate, but y rate, formaldehyde, oxalate, tartrate, pyruvate, malate, 
fumarate, succinate, a-ketoglutarate, citrate, irafls-aconitate, glu-
tamine, glutamate, asparagine, aspartate, and glycine. All were tested at 
0.01 M concentration. 
Starch Degradation 
In Chlamydomonas cells grown on a diurnal rhythm, starch is accumu-
lated within the chloroplast during the light phase and degraded in the 
dark. This cyclic starch metabolism of C. reinhardtii has been used as 
the basis for a theoretical model of storage material utilization in algae 
by Cohen and Parnas (1976). In continuous light either in the presence or 
absence of acetate, starch breakdown is greater under anaerobic (nitro-
gen) than aerobic conditions, a phenomenon known as the Pasteur effect 
(Peavey et al., 1983). Deposits which appear to be starch are seen sur-
rounding the pyrenoid, and cytochemical studies have shown that most 
of this material is amylase-sensitive (Hirschberg et al., 1981). In dark-
grown cells, starch granules are also seen scattered throughout the chlo-
roplast matrix (Ohad et al., 1967a). As in higher plants (Preiss, 1982), 
starch synthesis and degradation in algae appear to be regulated by the 
activities of ADP-glucose pyrophosphorylase and amylase, but in con-
trast to the situation in plants, in Chlamydomonas these enzymes appear 
to be associated with chloroplast fractions (Lev i and Gibbs, 1984). Both 
enzymes were observed to be present throughout the cell cycle but to 
reach peak activity about 4 hr into the dark phase in C. reinhardtii cells 
grown on a 12:12 light : dark cycle. Lev i and Gibbs partially purified and 
characterized an α-amylase from C. reinhardtii, similar to α-amylase 
from leaves, but did not eliminate the possibility that the alga also has a 
/3-amylase activity. They also identified activities of dextrinase, maltase, 
and hexokinase. The presence of hexokinase suggests that despite the 
inability of C. reinhardtii to grow on exogenous glucose, it is able to use 
glucose arising internally. Delvalle and Asensio (1978) identified A T P -
dependent hexose kinase activity for glucose and fructose, but not 
mannose. 
Starch Degradation 223 
The enzymes of glycolysisfrom fructose through 3-phosphoglycerate 
(Figure 6.1) may also be localized primarily in the chloroplast (Klein , 
1984, 1986). Kreuzberg and Martin (1984) and Gfeller and Gibbs (1984) 
inferred from their data that glycolysis probably predominates over the 
pentose phosphate pathway for sugar degradation in Chlamydomonas. 
The enzymes converting 3-phosphoglycerate to pyruvate are found out-
side the chloroplast fraction, as are malate dehydrogenase ( N A D
+
) and 
isocitrate dehydrogenase (Klein, 1984, 1986). A second malate dehydro-
genase activity, which is NADP
+
-dependent , shows a mixed distribu-
tion. This pattern of enzyme localization differs from that of higher 
plants but is seen in other green algae. T w o isozymic forms have been 
found for a number of additional enzymes of the glycolytic and oxidative 
pentose phosphate pathways, including aldolase (Guerrini et al., 1971a), 
glucose-6-phosphate isomerase, phosphoglucomutase, glucose-6-phos-
phate dehydrogenase, and 6-phosphoglucose dehydrogenase (Herbert et 
al., 1979). These may be separately allocated in chloroplast and cyto-
plasmic fractions (Klein, 1984). A similar division is seen in higher plants 
(Schnarrenberger et al., 1983). 
Under anaerobic conditions in the dark, starch fermentation in 
Chlamydomonas, Chlor ogonium, and Chlorella leads to production of 
formate, acetate, and ethanol (Klein and Betz, 1978a,b; Kreuzberg, 
1984a; Figure 6.1). Kreuzberg (1984a) and Gfeller and Gibbs (1984) con-
curred in finding that in C. reinhardtii these three compounds were 
produced in a 2 : 1 : 1 ratio, with small amounts of H 2 and C 0 2 also being 
starch 
I 
glucose-6-phosphate 
I 
pentose fructose-6-phosphate 
phosphate pathway? 
glyceraldehyde-3-phosphate 
I 
phosphoglycerate 
tricarboxylic acid 
cycle pyruvate • D-lactate 
\ / \ 
glyoxylate cycle •* acetyl CoA formate 
I \ 
acetaldehyde • acetate 
/ 
ethanol 
Figure 6.1 Schematic view of glycolysis in Chlamydomonas, adapted from Gfeller and 
Gibbs (1984) with permission of the American Society of Plant Physiologists. 
224 6. Metabolism 
released. In C. moewusii, large amounts of glycerol were produced 
(Klein and Betz, 1978a), whereas in C. reinhardtii glycerol and D-lactate 
were seen only in minor amounts and at low pH (Gfeller and Gibbs, 
1984; Kreuzberg, 1984a). Klein and Betz also reported substantial H 2 
release by C. moewusii cells. In C. reinhardtii cells in the light but with 
photosynthesis inhibited by D C M U [3,(3,4-dichlorophenyl-l,l-dimeth-
ylurea)], FCCP (carbonyl cyanide-p-trifluoromethoxyphenylhydrazone, 
an uncoupler of photophosphorylation), and the presence of the F-60 
mutation blocking photosynthetic carbon assimilation, acetate and for-
mate were produced in a 1 : 1 ratio, and ethanol production was inhibited 
(Gfeller and Gibbs, 1984). Kreuzberg (1984a) and Gfeller and Gibbs 
(1984) presented evidence for formate production from pyruvate medi-
ated by pyruvate formate lyase, a pathway previously thought to be 
restricted to prokaryotic organisms. This enzyme appears to be present 
even under aerobic conditions, and its activation on transfer to an anaer-
obic environment is not blocked by inhibitors of protein synthesis 
(Kreuzberg, 1984a). Phosphotransacetylase and acetate kinase activities 
were also detected. Lactate is oxidized rapidly under aerobic conditions 
in air-grown cells but is not metabolized in C0 2-grown cells ( D . W . 
Husic and Tolbert, 1985; D. W . Husic, personal communication). Fer-
mentation induced by transfer to anaerobic conditions showed an oscil-
latory pattern, as measured by ethanol and acetate production, with a 
mean period length of 59 min (Kreuzberg and Martin, 1984). Changes 
with the same period length in A T P , A D P , and A M P content were also 
observed. 
Kreuzberg et al. (1987) reported that pyruvate dehydrogenase, pyru-
vate formate lyase, and lactate dehydrogenase (pyruvate reductase) 
were found in both mitochondrial and chloroplast fractions, while phos-
photransacetylase and acetate kinase were primarily found in the mito-
chondrial fraction. Alcohol dehydrogenase was distributed between mi-
tochondria and cytoplasm. Aldehyde dehydrogenase was found both in 
the cytoplasm and in chloroplasts, and pyruvate decarboxylase was 
found only in the cytoplasm. 
Gfeller and Gibbs (1984) were able to construct a "balance sheet" for 
starch and its end products based on analysis of fermentative products 
under different experimental conditions (darkness or light and with or 
without D C M U and/or F C C P ) . T w o formate molecules are produced for 
each glucose consumed, and the total of acetate plus ethanol equals the 
formate. Light inhibits ethanol production under all conditions, and ace-
tate and formate are then produced in equimolar amounts. Whether 
acetate is formed directly from acetyl-CoA by a deacylase or is formed 
by acetaldehyde dehydrogenase has not been established (Figure 6.1). 
Very little lactate is formed in long-term (6-hr) experiments under 
anaerobic conditions. However , with pulse labeling during short-term 
(<30-min) experiments, D . W . Husic and Tolbert (1985) demonstrated 
that D-lactate accumulates rapidly. It appears to be oxidized slowly by 
Respiration 225 
the glycolate dehydrogenase associated with the mitochondrial mem-
brane (Nelson and Tolbert, 1970; Beezley et al., 1976). Husic and 
Tolbert showed that lactate was formed by reduction of pyruvate in an 
irreversible reaction catalyzed by pyruvate reductase (equivalent to the 
lactate dehydrogenase studied by Kreuzberg and colleagues). Since lac-
tate accumulates only during anaerobic conditions, Husic and Tolbert 
postulated that the enzyme is functioning to reoxidize N A D H . This 
enzyme is contrasted with a somewhat analogous metabolic step in 
higher plants, in which L-lactate rather than the D-isomer is formed 
under anaerobic conditions by an enzyme that catalyzes a reversible 
reaction. 
Respiration 
Difficulties in isolating purified mitochondria from Chlamydomonas 
have hampered physiological investigation of respiration, although ul-
trastructural studies of mitochondria have been fairly extensive (Schötz 
et al., 1972; Wiseman et al., 1977a; see also Chapter 3). The presence of 
at least some of the tricarboxylic acid cycle enzymes has been docu-
mented (see Table 6.1), and the presumption is that this cycle is func-
tional together with the glyoxylate cycle, as in other acetate flagellates 
(see Haigh and Bee vers, 1962; L loyd , 1974). Bothe and Nolteernsting 
(1975) reported that C. reinhardtii cells had high levels of lipoic acid, one 
of the cofactors of the pyruvate dehydrogenase complex, and concluded 
that the alga probably used this complex to form acetyl-CoA from pyru-
vate. The cyanobacterium Anabaena cylindrica, in contrast, was shown 
to accomplish this reaction by ferredoxin oxidoreductase, an enzyme 
previously described in anaerobic bacteria. 
Isocitrate dehydrogenase activity was reported in C. reinhardtii by 
Hess and Tolbert (1967a) and Ramaley and Hudock (1973), and in C. 
segnis by Foo and Badour (1977). Isocitrate lyase, the key enzyme of the 
glyoxylate cycle, is not detected in wild-type cells grown phototroph-
ically but is induced by exogenous acetate (Wiseman et al., 1977a). A 
mutant of C. dysosmos unable to grow on acetate in the dark was iso-
lated by Lewin (1954b) and its defect identified by Neilson et al. (1972) as 
failure to synthesize isocitrate lyase. Wiseman et al. (1977a) showed that 
phenotypically similar dark-dier (dk) mutants of C. reinhardtii could be 
separated into two classes according to their sensitivity to fluoroacetate, 
an inhibitor of aconitase (Kates and Jones, 1964b). Permeability and 
glyoxylate cycle mutants were not inhibited by this compound, but mu-
tants with defects elsewhere in respiration were sensitive (see be low) . 
Themitochondrial electron transport chain of Chlamydomonas re-
sembles that of plants in having a classical cyanide-sensitive pathway 
plus a cyanide-insensitive, salicylhydroxamic acid-sensitive branch (see 
Wiseman et al., 1977a). The relative activity of these two pathways may 
T a b l e 6.3 S u m m a r y of B iochemica l a n d Ul t rastructura l Proper t ies of N i n e M e n d e l i a n D a r k - D i e r M u t a n t s 
C o m p a r e d to Wi ld T y p e (dk )a 
Fluoro- Cyto- Antimycin or Whole cell Mitochondria 
acetate Acetate Isocitrate Cyanide- chrome rotenone sensitive ultrastructure 
sensi- incorpo- lyase sensitive oxidase NADH-cytochrome c (except mito- DAB 
Genotype tivity ration activity respiration activity reductase chondria) Ultrastructure staining 
dk+ Yes + + + + + Normal Normal + 
dk-32 Yes + + ± - - Normal Grossly altered -
dk-34 Yes + + - - Normal Grossly altered -
dk-52 No + + + Reduced electron Reduced electron -
density of density of 
membranes membranes 
dk-76 Yes + + ± ± ± Normal Normal -
dk-80 Yes ± + - - + Swollen endo- Swollen cristae -
plasmic reticu-
lum 
dk-97 Yes + - - + Normal Normal -
dk-105 Yes + + ± + Normal Normal + 
dk-110 Yes + - - + Normal Precipitated ± 
matrix material 
dk-148 Yes + + - ± + Normal Precipitated + 
matrix material 
α From Wiseman et al. (1977a). Reproduced from The Journal of Cell Biology, 1977, 53, 56-77 by copyright permission of The Rockefeller University Press. 
Respiration 227 
vary with cell physiology; Hommersand and Thimann (1965) reported a 
switch from cyanide-sensitive to -insensitive respiration during germina-
tion of zygospores, but this interesting observation has not been fol-
lowed up. Four of the obligate photoautotrophic mutants isolated by 
Wiseman et al. have very low levels of cyanide-sensitive respiration 
compared to wild-type Chlamydomonas and are deficient in cytochrome 
oxidase activity, measured either in cell extracts or cytochemically with 
diaminobenzidine (Table 6.3). Four other mutants have intermediate 
levels of cyanide-sensitive respiration. In three of the latter group, both 
antimycin- and rotenone-sensitive NADH-cytochrome c reductase ac-
tivities are lower than those observed in wild-type cells. Some, but not 
all, of the dk mutants have gross alterations in mitochondrial ultra-
structure. 
Kates and Jones (1964b) studied the effects on C. reinhardtii of block-
ing the tricarboxylic acid cycle with fluoroacetate on synthesis of other 
metabolic intermediates, particularly amino acids. Cells treated with 
fluoroacetate and given [
1 4
C ] 0 2 under conditions permitting photosyn-
thesis accumulated labeled citrate at very high levels, and synthesis of 
the other T C A cycle intermediates was inhibited. Very little label ap-
peared in either glutamate or aspartate compared to controls, but alanine 
and glutamine continued to be synthesized. This pattern of labeling is 
consistent with later studies on nitrogen assimilation and amino acid 
synthesis (see below) . 
Inability to grow in the dark on acetate is occasionally encountered 
spontaneously in stocks of C. reinhardtii selected for other reasons. 
Thompson et al. (1985) investigated this problem in a stock of the cell 
wall-deficient mutant cw-15. Death appeared to occur from swelling, 
presumably due to a malfunction in osmoregulation, after two or more 
cell divisions in the dark. Respiration in these cells appeared to be 
normal. Thompson et al. found that exposure of cells to dim blue light, 
insufficient for photosynthesis, prevented dark lethality, and proposed a 
model involving blue-light-induced synthesis of specific proteins neces-
sary for osmoregulation to account for this effect. Dark-tolerant strains 
were presumed to have constitutive synthesis of the postulated proteins. 
The Glycolate Pathway 
The oxygenase activity of ribulose bisphosphate carboxylase mediates 
the reaction of ribulose bisphosphate with molecular oxygen to form 
phosphoglycolate and 3-phosphoglycerate. The regulation with respect 
to C 0 2 concentration of this process, known as photorespiration, is 
discussed in more detail in Chapter 7. The reviews by Tolbert and col-
leagues (Tolbert, 1974, 1979; Husic et al., 1987) provide additional infor-
mation comparing photorespiration and glycolate metabolism in algae 
and higher plants. Phosphoglycolate is hydrolyzed by a specific phos-
phatase to release glycolate, which is excreted from the chloroplast and 
enters the C 2 pathway illustrated in Figure 6.2. Bruin et al. (1970) docu-
228 6. Metabolism 
ο 2 
ribulose bisphosphate • phosphoglycerate + phosphoglycolate 
[1] 
2 phosphoglycolate • 2 glycolate >-2 glyoxylate • 
[2] [3] [4] 
C 0 2 + N H 3 
2 glycine • serine • hydroxy pyruvate • 
[5,6] [7] [8] 
glycerate • phosphoglycerate 
[9] 
Enzymes: 
1 : ribulose bisphosphate carboxylase/oxygenase 
2: phosphoglycolate phosphatase 
3: glycolate dehydrogenase 
4: glutamate:glyoxylate aminotransferase 
5: glycine decarboxylase 
6: serine hydroxymethyltransferase 
7: serine:pyruvate transaminase 
8: hydroxy pyruvate reductase 
9: glycerate kinase 
Figure 6.2 Pathway of glycolate metabolism, adapted from Bruin et al. (1970). 
mented the existence of this pathway in Chlamydomonas and other 
green algae by tracing and comparing assimilation of exogenous [
1 4
C ] -
glycolate and photosynthetically fixed [
, 4
C ] 0 2 , and by demonstrating the 
presence of the participant enzymes of this pathway in cells grown on 
air. In contrast to higher plants, in which these enzymes are peroxi-
somal, in Chlamydomonas glycolate metabolism takes place in the mito-
chondria. Glycolate dehydrogenase is bound to the mitochondrial mem-
brane and is different from the peroxisomal glycolate oxidase of higher 
plants (Tolbert and Hess, 1966; Hess and Tolbert, 1967a; Nelson and 
Tolbert, 1969, 1970; Codd et al., 1969; Frederick et al., 1976; Stabenau, 
1974; Paul and Volcani, 1976; Beezley et al., 1976; Gruber and Fred-
erick, 1977). The transaminases and hydroxypyruvate reductase are 
also mitochondrial. The glycerate formed by hydroxypyruvate reduc-
tase is returned to the chloroplast and phosphorylated to form phos-
phoglycerate, which reenters photosynthetic carbon metabolism. 
The glycolate pathway is operative at low C 0 2 tension and is sup-
pressed by high C 0 2 or by inhibition of carbonic anhydrase (see Badger 
et al., 1980; Spencer and Togasaki, 1981). Under the latter conditions, 
Respiration 229 
glycolate is excreted into the medium (Tolbert and Zill, 1956; Nelson 
and Tolbert, 1969; Tolbert et al., 1983; Moroney et al., 1986a). Hess and 
Tolbert (1967b) found that glycolate was accumulated in cells grown in 
blue light (400-500 nm), but that malate, aspartate, glutamate, and 
alanine were the primary products of [
1 4
C ] 0 2 fixation by cells grown in 
red light (>600 nm). Mitochondrial respiration, that is, residual 0 2 up-
take, is maintained during photosynthesis under saturating C 0 2 , when 
the glycolate pathway should be maximally inhibited (Peltier and Thi-
bault, 1985a,b). At low C 0 2 tension, glycolate excretion is maximal in 
early G i phase cells and is minimal during cell division of C. reinhardtii 
grown synchronously on a 16:8 hr light : dark cycle (Kates and Jones, 
1966; Chang and Tolbert, 1970). The fraction of glycolate that is ex-
creted rather than metabolized is greatly stimulated by aminooxyacetate 
or aminoacetonitrile (Tolbert et al., 1983; Moroney et al., 1986a), which 
block the pathway at the glyoxylate-serine aminotransferase and gly-
cine-serine interconversion steps, respectively. This glycolate appears 
to result from R U B I S C O oxygenase activity. From these studies Moro-
ney et al. (1986a) estimated a rate of glycolate flux through the pathway 
in air-grown cells equivalent to 5-10 ^moles glycolate/hr per mg chloro-
phyll. 
D . W . Husic and Tolbert (1987b) further explored the connectionbetween respiration and glycolate metabolism in a mutant strain (dk-97) 
deficient in cytochrome oxidase activity. When the residual (cyanide-
insensitive) respiration was inhibited by salicylhydroxamic acid in the 
mutant under photosynthetic conditions, glycolate oxidation was 
blocked, and glycolate was accumulated and excreted. In wild-type 
cells, respiratory electron transport through cytochrome oxidase contin-
ued under these conditions, and glycolate excretion did not increase, 
suggesting that glycolate dehydrogenase activity is linked to mitochon-
drial electron transport. Oxidation of D-lactate appears to be similarly 
dependent on mitochondrial respiration (Husic and Tolbert, 1987b). Ex-
ogenous glycolate can be assimilated only in photosynthetically compe-
tent cells, and this assimilation occurs only at concentrations of glyco-
late substantially above the level likely to be found in nature (Spencer 
and Togasaki, 1981). Thus this is probably not an important function 
under natural conditions. King and Togasaki (1974) reported that high 
levels of exogenous glycolate inhibited growth of Chlamydomonas cells 
and proposed selection for glycolate resistance as a means of obtaining 
mutants blocked in glycolate metabolism. 
H . D . Husic and Tolbert (1984, 1985) have described the phosphogly-
colate phosphatase of C. reinhardtii. This enzyme resembles its counter-
part in higher plants and is sufficiently active to account for the observed 
rate of flux through the glycolate pathway. N A D H : hydroxypyruvate 
reductase and N A D P H : glyoxylate reductase have been characterized 
by D . W . Husic and Tolbert (1987a). 
230 6. Metabolism 
Chlororespiration 
Bennoun (1982) presented evidence that thylakoid membranes of 
Chlamydomonas after dark adaptation used molecular oxygen to oxidize 
photosynthetic plastoquinone, which could be reduced by N A D H using 
a thylakoid-bound iron-sulfur oxidoreductase (Godde, 1982; Gfeller and 
Gibbs, 1985). A n electrochemical gradient was shown to form across the 
thylakoid membrane as a result of this N A D H oxidation and by reverse 
functioning of chloroplast ATPase . Bennoun suggested that this process 
would recycle A T P and N A D ( P ) H generated by glycolysis. Mutants 
blocked in photosynthesis between plastoquinone and photosystem I 
still showed the chlororespiration phenomenon, suggesting that some 
other electron carriers might be involved in this process (Bennoun, 
1983). Subsequent studies by Kreuzberg (1984b), Gfeller and Gibbs 
(1985), and Maione and Gibbs (1986b) have confirmed the likelihood of a 
plastoquinone-mediated chloroplast oxygen consumption. Kreuzberg 
also identified alcohol dehydrogenase and lactate dehydrogenase activi-
ties in the chloroplast fraction which he suggested might be used in 
reoxidation of reducing equivalents. 
Hydrogenase 
Hydrogenase activity has been found in most species of Chlamydo-
monas examined (Table 6.4). Under anaerobic conditions in light and in 
the absence of C 0 2 , hydrogenases function to release H 2 and 0 2 from 
water (biophotolysis; see Bothe, 1982). They can also catalyze C 0 2 
reduction in algae illuminated after incubation with H 2 under anaerobic 
conditions (photoreduction) and can reduce various electron acceptors 
in the dark (see Kessler, 1974, for review; see Maione and Gibbs, 1986a, 
for a summary of more recent work) . Evolution of H 2 from anaerobic 
cells also occurs in the dark, particularly in C. moewusii (Healey, 1970b; 
Wang et al., 1971; Klein and Betz, 1978b), and appears to be coupled 
with starch degradation. In C. reinhardtii cells growing on a light : dark 
T a b l e 6.4 Chlamydomonas S p e c i e s in W h i c h H y d r o g e n a s e 
Activi ty H a s B e e n D e m o n s t r a t e d
3 
Species Reference 
C. debaryana Healey (1970a) 
C. dysosmos Healey (1970a) 
C. eugametos Abeles (1964); Ward (1970a) 
C. intermedia Ward (1970a) 
C. moewusii Frenkel (1949, 1951); Frenkel and Rieger (1951); Ward (1970a,b) 
C. reinhardtii Ben-Amotz et al. (1975); see text for additional citations 
a
 Modified from Kessler (1974). 
Respiration 231 
cycle, starch accumulation is seen in the light, and hydrogen evolution is 
observed throughout the dark phase if a very small amount of 0 2 is 
present (Miura et al., 1982). The biophotolysis reaction uses electrons 
from photosystem I and is stimulated by prolonged adaptation to anaero-
bic, I0W-CO2 conditions (Stuart and Gaffron, 1972a,b; Ben-Amotz 
and Gibbs, 1975; Zakrzhevskii et al., 1975, 1977; Oshchepkov et al., 
1978). Chlamydomonas reinhardtii has proved to be extremely toler-
ant of this stressful situation, suggesting that it may be a useful organ-
ism for investigation of potential industrial uses of the process (Green-
baum, 1982a,b; Greenbaum et al., 1983a; Reeves and Greenbaum, 
1985). 
Bamberger et al. (1982; see also Gibbs et al., 1986) measured release 
of C O 2 and H 2 under anaerobic conditions in the F-60 strain blocked in 
the photosynthetic carbon cycle. In the dark, addition of an uncoupler of 
phosphorylation increased starch breakdown and C 0 2 release but de-
pressed H 2 formation. In the light, acetate stimulated release of both 
C 0 2 and H 2 , but not starch breakdown; the uncoupler increased starch 
breakdown as well. Aparicio et al. (1985) found that hydrogen evolution 
was stimulated by provision of N H 4
+
 as sole nitrogen source, but inhib-
ited by N O 3 " or N 0 2 ~ . Maximal H 2 evolution was seen at low light 
intensity, and high light intensity inhibited the reaction. Synthesis of 
hydrogenase is seen within an hour of dark anaerobic adaptation and is 
inhibited by cycloheximide (Klein and Betz, 1978a; Yanyushin, 1979-
1982b; Matoura and Picaud, 1987). Addition of acetate accelerates the 
appearance of hydrogenase activity, but uncouplers prevent it, probably 
by interfering with an energy-requiring activation of the enzyme (Lien 
and San Pietro, 1981). A scheme coupling hydrogenase to photosyn-
thetic electron transport and chlororespiration via NADPH-plas to-
quinone oxidoreductase has been proposed by Maione and Gibbs 
(1986b). 
Ward (1970b) distinguished several isozymic forms of hydrogenase in 
C. moewusii cells and described their activation in cell-free extracts 
after A T P depletion. Hydrogenase from C. reinhardtii has been partially 
purified by Roessler and Lien (1982, 1984a-c). It is an iron-containing 
acidic enzyme of about 45 kDa, and is generally similar to bacterial 
hydrogenases, although it is less stable than most of these at high tem-
perature (55°C). Activity of the enzyme in vitro with methyl viologen is 
stimulated by various anions, especially iodide, thiocyanate, and bro-
mide (Roessler and Lien, 1982). With ferredoxin, the natural electron 
mediator, no anion stimulation is seen in vitro. Roessler and Lien 
(1984c) interpreted these results to suggest a positively charged region 
near the catalytic site of the enzyme, very likely containing lysine and/or 
arginine residues. The enzyme is irreversibly inactivated by oxygen and 
is reversibly inhibited by C 0 2 (Erbes et al., 1979; Maione and Gibbs, 
1986a). 
232 6. Metabolism 
Lipid Metabolism 
Photo- and mixotrophically grown cells of C. reinhardtii contain on the 
order of 18-24 mg of ether-soluble lipid per gram of dry weight (~10
9 
cells); heterotrophically grown cells contain less, about 8-12 mg (Ei-
chenberger, 1976). Of this total, chlorophyll accounts for about 19% in 
mixotrophically grown cells, monogalactosyl diglycerides ( M G D G ) 
about 26%, and digalactosyl diglycerides ( D G D G ) another 19%. Phos-
phatidyl ethanolamine, phosphatidyl glycerol, phosphatidyl choline, and 
sulfolipid together account for about 14% of cellular polar lipids (Eichen-
berger, 1976; Janero and Barrnett, 1981c,d). The remaining ether-soluble 
lipid, 24-30% of the total, comprises carotenoids, sterols, and severalother components. Prominent among the latter is an unusual membrane 
lipid, diacylglyceryl trimethyl homoserine ( D G T S ) . A high level of this 
compound is associated with relatively low levels of phosphatidyl cho-
line in Chlamydomonas and in the Chrysophyte alga Ochromonas 
danica. Some other algae, including Chlorella, Eu g le na, and Fucus, 
have high levels of phosphatidyl choline but low DGTS (Eichenberger 
and Boschetti, 1978; Eichenberger, 1982; Janero and Barrnett, 1982e). 
( D G T S was formerly called "lipid A " in Ochromonas and "lipid X " in 
Chlamydomonas.) Chlamydomonas reinhardtii cells incorporate labeled 
oleic acid into D G T S and subsequently desaturate it to linoleic and y-
linolenic acids. Schlapfer and Eichenberger (1983) inferred from these 
observations that DGTS may function primarily as an acyl carrier for 
desaturation of oleic and linoleic acids. This function is served by phos-
phatidyl choline in higher plants. 
Cellular lipids of Chlamydomonas show marked partitioning between 
chloroplast and cytoplasmic compartments. Glycolipids account for 70-
80% of the thylakoid membrane lipid content, with DGTS and phos-
phatidyl glycerol in approximately equal proportions making up the re-
mainder (Eichenberger et al., 1977; Janero and Barrnett, 1981b, 1982e,f). 
Phosphatidyl ethanolamine and phosphatidyl choline are not found in 
the association with thylakoid membranes, whereas more than 50% of 
total cellular phosphatidyl glycerol is chloroplast-associated (Janero and 
Barrnett, 1981c; Mendiola-Morgenthaler et al., 1985b). Diphosphatidyl 
glycerol (cardiolipin) is specifically associated with the mitochondrial 
inner membrane in higher plant cells. Janero and Barrnett (1982d) identi-
fied cardiolipin in C. reinhardtii cells and showed that its fatty acid 
composition was similar to that of the cell as a whole and to the thyla-
koid membrane, but they did not localize it to a specific cellular fraction. 
Glycerol-3-phosphate acyltransferase and lysophosphatidate acyltrans-
ferase activities are localized in the chloroplast envelope, in association 
with the thylakoid membranes, and in pyrenoid tubules (Jelsema et al., 
1982; Michaels et al., 1983). Galactosyl transferase is associated specifi-
cally with the chloroplast envelope (Mendiola-Morgenthaler et al., 
1985b). Lysophosphatidate acyltransferase activity is also seen near 
Lipid Metabolism 233 
the outer mitochondrial membrane in cytochemical studies. Hoppe and 
Schwenn (1981) reported association of sulfolipid synthesis with thyla-
koid membranes of Chlamydomonas, as is the case in higher plants. 
C i 6 and C\% fatty acids predominate in Chlamydomonas lipids (see 
Erwin, 1973; Gealt et al., 1981; Janero and Barrnett, 1981c). The thyla-
koid M G D G s contain mostly unsaturated C i 8 acids, whereas the DGDGs 
and sulfolipids contain a much higher proportion of saturated C i 6 acid 
(Janero and Barrnett, 1981b). Slight differences in relative proportions 
of fatty acids are seen when thylakoid membrane fractions are compared 
to whole cell preparations, but these differences are much less dramatic 
than those among M G D G , D G D G , and sulfolipids. 2-Hydroxy fatty 
acids are also seen. The most abundant of these is 2-hydroxyhexadeca-
noic acid but longer-chain 2-hydroxy acids are also found, including 
substantial amounts of the C 26 compound 2-hydroxyhexacosanoic acid 
(Matsumoto et al., 1984). The corresponding saturated and unsaturated 
normal C 2 6 acids are not seen. 
The predominant sterols in wild-type C. reinhardtii cells and flagella 
are ergosterol and 7-dehydroporiferasterol (Patterson, 1974; Gealt et al., 
1981). Bard et al. (1978) also reported finding smaller amounts of several 
additional sterols. Three mutants isolated by Bard et al. as nystatin-
resistant proved to have altered sterol composition. One (KD7) ap-
peared unable to reduce the C-25 double bond required for ergosterol 
synthesis; the other two (KD 16 and KD21) appeared to lack the 22(23) 
desaturase activity. Sterol distribution in the plasma membrane has been 
studied with freeze-fracture and cytochemical markers by Robenek and 
Melkonian (1981). 
Beck and Levine (1977) and Janero and Barrnett (1981a, 1982a-c) 
have followed synthesis of thylakoid membrane lipids, pigment, and 
sterols over the cell cycle in synchronously grown cells. All components 
were maximally synthesized in the light period (mid- to late G j ) , but 
distinct temporal variations were seen among individual compounds, 
indicative of a multistep assembly process. Changes in thylakoid mem-
brane stucture and function, particularly of photosystem I I , after lipase 
treatment were described by Okayama et al. (1971). 
Sirevâg and Levine (1972) identified fatty acid synthetase activity in 
extracts of C. reinhardtii cells. In contrast to Euglena, in which two 
distinct fatty acid synthetase activities were reported (Delo et al., 1971), 
only a single activity, dependent on added acyl carrier protein, was 
detected in Chlamydomonas. In synchronously grown cells, the major 
products formed from acetyl-CoA and malonyl-CoA were palmitate 
( C ! 6) , stéarate ( C i 8) , and arachidate ( C 2 0) in the light period of growth, 
and predominantly shorter-chain fatty acids in the dark. Neither rifampi-
cin nor cycloheximide inhibited activity of fatty acid synthetase in these 
cultures, but spectinomycin, an inhibitor of organelle protein synthesis, 
reduced its activity significantly. Sirevâg and Levine concluded that 
fatty acid synthetase might therefore be synthesized on chloroplast ribo-
234 6. Metabolism 
somes, but they interpreted the rifampicin insensitivity to suggest that 
transcription might take place from a nuclear gene. Subsequent studies 
on the chloroplast genetic system have not documented any cases of 
chloroplast translation of nuclear m R N A s (see Chapter 8), and despite 
the extensive subsequent work on the chloroplast genome of Chlamy-
domonas, no evidence for a chloroplast-encoded fatty acid synthetase 
gene has so far been presented. Possibly the chloroplast component 
whose synthesis is spectinomycin-sensitive is a processing enzyme or 
membrane protein needed for assembly or binding of the fatty acid syn-
thetase. Further study on this problem is probably warranted. 
Nonspecific Phosphatases 
Five distinct phosphatase activities have been identified in C. reinhardtii 
cells: there are two constitutive acid phosphatases (one soluble and one 
particle-bound), derepressible soluble and bound alkaline phosphatases, 
and a derepressible neutral phosphatase active at a wide range of pH 
values (Guerrini et al., 1971b; Lien and Knutsen, 1972, 1973a; Matagne 
and Loppes, 1975; Nagy et al., 1981). Acid and alkaline phosphatases 
from Chlamydomonas acidophila have been described by Boavida and 
Heath (1986). The derepressible phosphatases of C. reinhardtii are in-
duced by removing inorganic phosphate from the culture medium and by 
addition of substrates such as /^-glycerophosphate. Induction of alkaline 
phosphatase is more rapid in acetate-containing medium than in minimal 
medium (Guerrini et al., 1971b). 
The two acid phosphatases are clearly distinct and are probably coded 
by different genes, since they are separately affected by nonallelic muta-
tions (Loppes and Matagne, 1973). Nagy et al. (1981) identified four 
isozymic forms of the soluble enzyme, which they postulated might 
result from posttranslational modification of a single gene product. 
Loppes and Matagne (1973) developed a colony assay for acid phospha-
tase activity based on hydrolysis of a-naphthylphosphate to release a-
naphthol, which is coupled to a diazonium salt to produce a red color. 
Using this assay, they were able to identify two classes of mutants (P2 
and Pa), each deficient in one of the acid phosphatase activities. A 
double mutant strain deficient in bothactivities was then used to select 
mutants (PD) deficient in the neutral phosphatase (Matagne and 
Loppes , 1975). None of 10 mutants representing three unlinked PD loci 
made protein antigenically related to the wild-type neutral phosphatase, 
nor did a temperature-sensitive mutant at one of these loci (Loppes et 
al., 1977; Loppes, 1977a). Working on the assumption that none of these 
loci was the structural gene for the enzyme, Loppes (1978) then at-
tempted to isolate a mutant producing a heat-sensitive phosphatase. 
Although such a mutant was indeed found (PDS), further studies showed 
that its phosphatase could be modified to the wild-type form by extracts 
Nitrogen Assimilation 235 
of wild-type cells, suggesting that the mutation isolated was in a gene 
involved in posttranslational modification of the phosphatase. T o date, 
no mutations have been confirmed to be in structural genes for the 
alkaline or neutral phosphatases. 
Disintegration of the mother cell wall after division of wild-type C. 
reinhardtii releases phosphatases into the culture medium (Lien and 
Knutsen, 1973a). Cells of the wall-deficient mutant cw-15 also release 
phosphatase activity (Loppes, 1976a). Matagne et al. (1976a,b) found by 
cytochemical staining that the soluble acid phosphatase was located 
primarily in vacuoles, while the neutral phosphatase was found in vacu-
oles and in the periplasmic space beneath the cell wall. The insoluble 
acid phosphatase was associated with cellular debris but could not be 
visualized cytochemically, nor were the alkaline phosphatases detected 
by these means. Indirect evidence from Loppes and Deltour (1975) sug-
gests that the alkaline phosphatases may also be located near the cell 
wall: attempting to isolate mutants deficient in these enzymes, they 
selected colonies deficient in all phosphatase activities from a muta-
genized culture of a triple mutant (P2PaPD4) lacking the acid and neutral 
phosphatases. Al l the mutants obtained proved to be cell wall-deficient 
strains that leaked phosphatase into the medium. Later studies (Loppes 
and Deltour, 1978, 1981) led to isolation of two temperature-sensitive 
cell wall mutants, one with normal phosphatase activities but showing 
leakage of phosphatase into the medium, and the other with altered 
neutral and alkaline phosphatase activities. The relation of the cell wall 
and phosphatase defects in the latter mutant are unclear. 
Nitrogen Assimilation 
Most algae use N H 4
+
 in preference to N 0 3 ~ as a nitrogen source, that is, 
if both ions are present, N 0 3 ~ will not be utilized until the N H 4
+
 is 
exhausted (Syrett, 1962; Thacker and Syrett, 1972a). A few species of 
Chlamydomonas have been reported to show no preferential N H 4
+
 utili-
zation (Cain, 1965). Except for the Ebersold-Levine strain of C. 
reinhardtii, however, which has natural mutations blocking nitrate re-
ductase activity (see be low) , most Chlamydomonas species are in any 
case capable of assimilating N 0 3 ~ , N 0 2 ~ , and N H 4
+
 as sole nitrogen 
sources, as well as any of a number of other compounds (Table 6.5). 
Chlamydomonas reinhardtii will also grow on urea, uric acid, aceta-
mide, glutamine, ornithine, arginine, hypoxanthine, allantoin, allantoic 
acid, guanine, and adenine (Sager and Granick, 1953; Cain, 1965; Gre-
sshoff, 1981a; Pineda et al., 1984a). N o species in Cain's survey was able 
to use cytosine, thymine, or uracil as its sole nitrogen source. 
T w o isolates of one species, C. Pulsatilla, cannot use either nitrate or 
ammonium but rather require an organic nitrogen source (Droop, 1961). 
A strain isolated from Finland ( C C A P 11/44) could use arginine, histi-
T a b l e 6.5 N i t rogen Sources Uti l ized by Chlamydomonas S p e c i e s
3 
Compounds utilized 
Species Urea Uric acid Acetamide Succinamide Adenine 
C. actinochloris - - - + + 
C. calyptrata
h 
+ + - + 
C. chlamydogama - - - -
C. carrosa + - + - + 
C. eugametos + - - - -
C. gloeogama - + - + 
C. gloeopara + + + + + 
C. inflexa + + + - + 
C. kakosmos + - + 
C. mexicana + - + + 
C. microsphaera v. acuta + + + - + 
C. microsphaerella + - + 
C. minuta + + + + 
C. moewusii + - - - + 
C. moewusii v. rotunda + + - - + 
C. mutabilis + - - - + 
C. peterfii + - + - -
C. radiata - - - - -
C. reinhardtii + + -
C. sectilis - - - - -
C. typhlos + + + - + 
Amino acids 
Ala Asn Glu Gin Gly Lys Orn Ser 
C. actinochloris + + - + + - + 
C. calyptrata
b - - - - - - -
C. chlamydogama - - + - - -
C. carrosa + + c - - -
C. eugametos - - - - - - -
C. gloeogama - - - - - + -
C. gloeopara - -h + + + -
C. inflexa - - + + - + + 
C. kakosmos - - - + - - -
C. mexicana - - - + -
C. microsphaera v. acuta + - - - - - -
C. microsphaerella - - - - - - -
C. minuta - - - + - - -
C. moewusii - + - - - + -
C. moewusii v. rotunda - + - - + - - -
C. mutabilis + - + + + + + 
C. peterfii - - - + - - -
C. radiata - - - - - - -
C. reinhardtii - - - - + -
C. sectilis - - - - - + -
C. typhlos - - - - - - — 
" Information primarily from Cain (1965). 
h
 Same as C. latifrons (Ettl, 1976a). 
' Some variation among different isolates of this species. 
''Cain (1965) stated that the UTEX 89 and 90 strains of C. reinhardtii did not grow on glutamine. 
However, the Ebersold-Levine wild-type strain appears to grow quite well using this compound as its 
sole nitrogen source. 
Nitrogen Assimilation 237 
dine, and lysine, while an isolate from Scotland used alanine but not 
lysine. Both isolates were obligate mixotrophs, requiring acetate or py-
ruvate for growth in the light and being unable to grow in the dark, and 
both required vitamin B i 2 . Hellebust and L e Gresley (1985) have found 
that an isolate of C. Pulsatilla from N e w Brunswick can grow hetero-
trophically on acetate and can use ammonium and several amino acids, 
but not nitrate or urea, as nitrogen sources. In nature C. Pulsatilla is 
found in rock pools below the nesting sites of sea birds, and its extensive 
organic needs are provided by the bird droppings. 
Uptake and assimilation of exogenous nitrate have been studied most 
thoroughly in Sager's wild-type strain of C. reinhardtii (21 gr mt
+
 and 
6145c y-1 mt~). Nitrate uptake is an energy-requiring process, stimu-
lated by nitrogen starvation and repressed by N H 4
+
 and by N 0 2
_
. In the 
absence of a carbon source (light plus C 0 2 , or acetate), neither N 0 3 ~ 
nor N H 4
+
 is assimilated, and cells in which photosynthesis is blocked by 
D C M U assimilate nitrogen only if acetate is added (Thacker and Syrett, 
1972a). Uptake of exogenous nitrite takes place by a permease-mediated 
system without a diffusion component that is distinguishable from the 
enzymatic nitrate reduction. The permease has an active site for nitrite 
that is not usable for nitrate transport (Cordoba et al., 1986). 
Thacker and Syrett (1972b) found that when cells grown on N H 4
+ 
were transferred to N 0 3 ~ , synthesis of nitrate reductase was dere-
pressed, with maximal activity being reached in 5-6 hr. Appearance of 
the enzyme is blocked by cycloheximide and by tungstate (Hipkin et al., 
1980). Addition of tungstate or N H 4
+
 to induced cultures also causes 
disappearance of existing nitrate reductase activity; nitrate protects the 
enzyme from degradation. Losada et al. (1973) suggested that N H 4
+ 
indirectly brought about reduction of the enzyme, possibly by uncou-
pling noncyclic photophosphorylation and thereby raising the level of 
reducing power in the cell. Treatment of C. reinhardtii cells in nitrogen-
free medium with methionine sulfoximine, an inhibitor of glutamine syn-
thetase, also inhibits nitrate reductase and produces excretion of N H 3 
into the medium, probably as a result of protein degradation (Hipkin et 
al., 1982; Florencio and Vega, 1983a). In cells grown on N 0 3 ~ , 
methionine sulfoximine inhibits N 0 3 ~ utilization, but not that of nitrite, 
and thus N H 3 isproduced in light by N 0 2 ~ reduction. However , in 
nitrogen-starved cells treated with methionine sulfoximine in the pres-
ence of nitrate, ammonium is excreted into the medium after nitrate 
reduction (Florencio and Vega, 1983a). 
N H 4
+
 represses nitrate reductase synthesis, but glutamine and gluta-
mate, the first organic compounds in the nitrogen assimilation pathway 
(see be low) , do not (Florencio and Vega, 1983c). Methylammonium also 
represses nitrate reductase synthesis. This compound is converted by 
glutamine synthetase to iV-methyl glutamine, which accumulates in the 
cells and does not seem to be further metabolized (Franco et al., 1984a). 
Exhaustion of methylammonium from the medium leads to derepression 
238 6. Metabolism 
of nitrate reductase synthesis concomitant with N-methyl glutamine ac-
cumulation in the cells. Franco et al. (1984a) concluded from these ex-
periments that N H 4
+
, not any of its metabolic products, is the natural 
corepressor of nitrate reductase synthesis. A methylammonium resis-
tant mutant (ma-1) defective in N H 3 and methylammonium uptake 
showed low intracellular levels of N H 3 and derepressed nitrate reduc-
tase activity in ammonium medium (Franco et al., 1987). 
A t low (atmospheric) C 0 2 levels in minimal medium, Chlamydomonas 
cells growing on N 0 3 ~ excrete Ν 0 2 ~ and N H 4
+
 into the culture medium. 
This process is promoted by blue light, which activates nitrate reduc-
tase, probably by a flavin-mediated reaction. Photosynthetically pro-
duced reductant is required. If the C 0 2 concentration is raised to 2%, 
excretion stops, and previously excreted N 0 2 " and N H 4
+
 are assimi-
lated (Azuara and Aparicio, 1983, 1984, 1985; Aparicio and Azuara, 
1984). 
The pentose phosphate pathway enzymes glucose-6-phosphate dehy-
drogenase and 6-phosphogluconate dehydrogenase are present at higher 
levels of activity in cells grown on nitrate than on ammonium medium 
and are greatly overproduced in all media in cells carrying the nitA 
mutation, which blocks nitrate reductase activity (Hipkin and Cannons, 
1985). These findings suggest coordinate regulation of nitrate assimila-
tion and pentose phosphate metabolism. 
Nitrate Reductase 
Nitrate reductase [ N A D ( P ) H : nitrate oxidoreductase] appears to be lo-
cated primarily in the pyrenoid of green algae (Lopez-Ruiz et al., 1985). 
This heteromultimeric complex has two separable activities (see Barea 
and Cardenas, 1975; Sosa and Cardenas, 1977; Franco et al., 1984b). A 
diaphorase is capable of mediating electron transfer in vitro from N A D H 
or N A D P H to oxidized acceptors such as cytochrome c, 2,6-di-
chlorophenolindophenol, potassium ferricyanide, or menadione. In vivo 
this reducing power is transferred to the terminal nitrate reductase to 
produce nitrite. The terminal reductase can be assayed in vitro with 
artificial electron donors such as reduced F A D , F M N , or viologens. The 
diaphorase, or NAD(P)H-cytochrome c reductase subunit, is a protein 
of about 45 kDa, and is associated with F A D and cytochrome b 5 57 
(Fernandez and Cardenas, 1983a,b; Franco et al., 1984b). The terminal 
reductase subunit appears to be equivalent to a protein of 67 kDa that is 
released by limited trypsin digestion of the native nitrate reductase com-
plex (Franco et al., 1984b). The diaphorase is active by itself, but the 
terminal subunit is not. Genetic evidence suggests that both subunits are 
coded by the nit-1 locus, which may therefore be dicistronic (Fernandez 
and Cardenas, 1982a, 1983c; Fernandez and Matagne, 1984, 1986). Stud-
ies with diploid cells formed from parents carrying different nit-1 muta-
tions indicate that the subunits are exchangeable to form hybrid en-
zymes (Fernandez and Matagne, 1986). Like the nitrate reductase of 
Nitrogen Assimilation 239 
fungi and higher plants, the complex includes a molybdenum-containing 
pterin cofactor (see Johnson, 1980). Complementation studies in vitro 
using cell-free preparations from nitrate reductase-deficient strains sug-
gest that in Chlamydomonas this cofactor must assemble with the pro-
tein subunit bearing the terminal reductase activity before the latter can 
interact with the diaphorase subunit (Fernandez and Cârdenas, 1981b). 
The native enzyme isolated by Franco et al. (1984b) had a mass of 220 
kDa and was presumed to consist of two each of the diaphorase and 
terminal subunits. Using the nitA (nit-la) mutant, Hipkin et al. (1985) 
found that enzyme extracted from cultures incubated in N 0 3 ~ medium 
was a single 390-kDa species, but enzyme from cultures incubated in 
nitrogen-free medium contained a 52-kDa protein with terminal reduc-
tase activity as well as several very large nitrate reductase complexes 
(225, 480 and 500 kDa). Wild-type cells on N 0 3 " produced a single 
enzyme species of 188 kDa. One concludes that the enzyme may exist in 
several forms. 
Mutants deficient in nitrate reductase activity are often insensitive to 
chlorate (Stouthamer, 1976; Cove , 1976a; Müller and Gräfe, 1978; Sosa 
et al., 1978; Nichols and Syrett, 1978). The simplest explanation for this 
effect is that wild-type nitrate reductase reduces chlorate to chlorite, 
which is toxic. However , not all chlorate-resistant mutants lack nitrate 
reductase (Nichols and Syrett, 1978), and mutants deficient in nitrate 
reductase may not necessarily be resistant to chlorate (see Cove , 1976b). 
Nichols and Syrett (1978) found that the largest group of chlorate-resis-
tant mutants arising from their experiments were able to grow on nitrate 
in the absence of acetate but not in its presence, but they were unable to 
explain this result. 
Chlamydomonas mutants have been found that lack the terminal re-
ductase but have diaphorase activity, that lack only the diaphorase, and 
that lack both activities (Table 6.6). Some of the mutants deficient in the 
terminal reductase also lack xanthine dehydrogenase and are unable to 
use hypoxanthine as a nitrogen source. Extracts from these mutants 
cannot restore nitrate reductase activity when combined with extracts of 
the Neurospora crassa mutant nit-1, whereas mutants deficient in termi-
nal reductase but not xanthine dehydrogenase can complement Neuro-
spora nit-1 in vitro. Fernandez and Cârdenas (1981a) concluded that the 
molybdenum cofactor of Chlamydomonas nitrate reductase is shared by 
xanthine dehydrogenase, as is also true in fungi and higher plants (Pate-
man et al., 1964; Κ . Y . L e e et al., 1974; Mendel and Müller, 1976). Like 
nitrate reductase, xanthine dehydrogenase activity is repressed in wild-
type cells grown on N H 4
+
; however, the molybdenum cofactor is still 
produced (Fernandez and Cârdenas, 1981a). Mutants lacking xanthine 
dehydrogenase do not appear to be at any growth disadvantage. In wild-
type cells grown on urea or aspartate as sole nitrogen source, very little 
nitrate reductase is synthesized, but xanthine dehydrogenase is still 
made at substantial levels. Thus these two enzymes are regulated sepa-
240 6. Metabolism 
Terminal Molybdenum Analogous mutants 
Strain'' Genotype Diaphorase reductase cofactor in other organisms 
305, nit A nit-1 α + + Neurospora nit-3 
Aspergillus nia D 10 
Tobacco nia-95 
Barley Az. 38 
301 nit-lb + - + No equivalent 
203, nitB nit-2 - - + Tobacco niai 
Ebersold-Levine 
wild type nit-1 nit-2 - - + — 
307 nit-3 + — Neurospora nit-1 
Aspergillus cnx 
104 nit-4 + - - Tobacco cnx 
21 gr nit-5 + + + — 
I 3 
nit-6 + + + — 
102 nit-5 nit-6 + - - — 
" Modified from Fernandez and Matagne (1984), with information on analogous mutants from Fernandez and Cardenas (1982a). 
For additional references, see nit entries in Chapter 11. 
* Numbered strains are those isolated by Sosa et al. (1978) 
rately (Fernandez and Cardenas, 1981a). Fernandez and Aguilar (1987) 
have characterized mutants deficient in the molybdenum cofactor. 
Theterminal nitrate reductase can be reversibly inactivated in vitro by 
reduction with K C N or dithionite and reactivated by oxidation with 
ferricyanide (see Cordoba et al., 1985). Nitrate protects the enzyme 
against inactivation (Barea et al., 1976). Added N A D H or N A D P H can 
also effect the reduction in extracts of wild-type cells, but not with 
enzyme from the diaphorase-deficient mutant 305 (nit-la). In vivo, ei-
ther nitrate deprivation or N H 3 addition leads to a comparable inactiva-
tion (Herrera et al., 1972; Florencio and Vega, 1982), which can be 
mediated by reductants other than N A D ( P ) H . In wild-type cells, N H 3 
blocks N 0 3 ~ uptake, but in the diaphorase-deficient mutant, N 0 3 ~ up-
take appears to be deregulated, and sufficient N 0 3 ~ enters the cells to 
reverse inactivation of the terminal reductase. The conclusion is that 
nitrate assimilation in Chlamydomonas is controlled at two levels, the 
uptake system and the activity of nitrate reductase itself, which is pro-
tected when N 0 3 " is available (Cordoba et al., 1985). Nitrate reductase 
can also undergo an irreversible inactivation in cells subjected to condi-
tions causing redox interconversion of the enzyme complex (Fernandez 
et al., 1986). This irreversible inactivating system seems to act preferen-
tially on the reversibly inactivated nitrate reductase, thus changing the 
enzyme turnover rate. 
Reduction of nitrite is accomplished by a ferredoxin nitrite reductase 
(Barea and Cardenas, 1975; Florencio and Vega, 1982, 1983a). Barea 
and Cardenas (1975) estimated the mass of this enzyme as 67 kDa and 
T a b l e 6.6 Ni t ra te R e d u c t a s e Muta t ions of C. reinhardtii
9 
Nitrogen Assimilation 241 
reported that it appeared to resemble nitrite reductases from Chlorella, 
Anabaena, and higher plants. 
Reduced nitrogen is incorporated into organic compounds primarily, 
possibly exclusively, by the glutamine synthetase-glutamate synthase 
system (Figure 6.3). Glutamine synthetase appears to be the rate-limit-
ing enzyme in nitrogen assimilation by Chlamydomonas (Cullimore and 
Sims, 1981b). This enzyme is deactivated by the glutamine analog 
methionine sulfoximine (Cullimore and Sims, 1980; Hipkin et al., 1982; 
Peltier and Thibault, 1983). Since methionine sulfoximine treatment of 
wild-type cells relieves NH 3-mediated repression of N 0 3 ~ uptake and 
inactivation of nitrate reductase (see above) , Cullimore and Sims (1981c) 
postulated that reversible deactivation of glutamine synthetase might 
regulate N 0 3 " assimilation. Transfer of N 0 3" - g r o w n cells to darkness 
and N H 4
+
 deactivates the enzyme; light restores activation (Cullimore, 
1981). Reactivation is partially inhibited by D C M U but is not affected by 
cycloheximide or chloramphenicol (Cullimore, 1981). Treatment with 
sulfhydryl reagents activates the enzyme in vitro. 
Florencio and Vega (1983b) separated two glutamine synthetase iso-
zymes, GSi and G S 2 , from C. reinhardtii cells grown on N 0 3 ~ . Both 
enzymes are large proteins (380 and 373 kDa, respectively), composed 
of eight subunits each. The G S 2 protein has an appreciably higher Km for 
N H 3 than does GSj and predominates in light-grown cells. In dark-
grown cells, GS | increases greatly and G S 2 becomes negligible. Both 
Enzymes: 
1 : nitrate reductase 
2: nitrite reductase 
3: glutamine synthetase 
4: glutamate synthase 
A: urea carboxylase 
B: allophanate lyase 
The Glutamine-Glutamate Cycle 
glutamate α-ketoglutarate 
2 glutamate 
urea + H C 0 3" — • a l l o p h a n a t e — + 2 N H 4
+
 + 2 H C 0 3" 
Figure 6.3 Pathways of nitrogen assimilation. 
242 6. Metabolism 
enzymes resemble the plant type of glutamine synthetase rather than the 
form found in photosynthetic prokaryotes. Very likely G S 2 is a chloro-
plast enzyme and is coupled to ferredoxin-glutamate synthase (see be-
l o w ) , while GSi is found outside the chloroplast fraction and is coupled 
to NADH-glutamate synthase. 
Glutamate synthase ( G O G A T ) is the second enzyme of the glutamine-
glutamate cycle (Figure 6.3). T w o distinct enzymes are present in C. 
reinhardtii (Cullimore and Sims, 1981a; Gal van et al., 1984; Marquez et 
al., 1984, 1986a). The NADH-specif ic G O G A T complex, presumed to be 
cytoplasmic, has two assayable activities, an N A D H diaphorase reduc-
ing ferricyanide, and methyl v io logen-GOGAT. It is specific for N A D H , 
in contrast to the higher plant equivalent, which is also active with 
N A D P H . N A D H - G O G A T of Chlamydomonas probably functions 
mainly in assimilation of exogenous N H 3 . Ferredoxin-GOGAT is a chlo-
roplast enzyme, which with its associated glutamine synthetase is proba-
bly primarily involved in reassimilation of N H 3 generated by photores-
piration (Marquez et al., 1986b). This protein is a single polypeptide 
chain of 146 kDa containing one F A D , one F M N , and one iron-sulfur 
cluster (Galvan et al., 1984; Marquez et al., 1986b). The N A D H gluta-
mate synthase is cold-labile and is inhibited by 0 2 . Its primary function 
appears to be in assimilation of N H 3 in the dark and in recycling of N H 3 
released from protein degradation (Marquez et al., 1986a). Both N A D H -
and ferredoxin-GOGAT activities are inhibited by the glutamine ana-
logue azaserine. 
The relationships between photorespiration, nitrogen assimilation, 
and protein catabolism have been investigated in Chlamydomonas by 
Cullimore and Sims (1980, 1981a-c), by Hipkin et al. (1982), and by 
Peltier and Thibault (1983, 1984). The glycolate pathway (see Figure 6.2) 
leads to production of N H 3 and C 0 2 from the conversion of glycine to 
serine, and this N H 3 is assimilated by the chloroplast GS 2- fe r redoxin-
G O G A T reactions. Cullimore and Sims (1980) showed that much of the 
nitrogen released in photorespiration is derived originally from protein 
catabolism rather than from newly synthesized glutamate. Hipkin et al. 
(1982) confirmed these results and further suggested that nitrogen star-
vation increased proteolysis, leading to release of ammonium in both 
light and darkness, which was readily observed in cultures treated with 
methionine sulfoximine to block glutamine synthetase activity. The 
reassimilation of nitrogen released by proteolysis may be important in 
synthesis of new proteins in gametogenesis (see Jones, 1970; Necas and 
Tetik, 1985). 
Although glutamate dehydrogenase has been demonstrated in C. 
reinhardtii, this enzyme probably functions only in catabolism and is not 
involved in N H 3 assimilation under normal circumstances (Cullimore 
and Sims, 1981b). However , when glutamine synthetase activity is 
blocked with methionine sulfoximine, some N H 3 uptake is still seen if 
the dissolved C 0 2 concentration of the medium is high (Hipkin et al., 
Nitrogen Assimilation 243 
1982), and this uptake is thought to be mediated by glutamate dehydro-
genase (Peltier and Thibault, 1983). Peltier and Thibault suggested that 
this process could account for about 30% of observed N H 3 assimilation 
under conditions of C 0 2 saturation. T w o distinct glutamate dehydro-
genase enzymes were found in C. reinhardtii by Kiv ic et al. (1969); one 
appeared to be chloroplast-associated. Both were active with either 
N A D or N A D P . 
Nearly all nitrogen assimilation work in Chlamydomonas has been 
done with C. reinhardtii. One exception is the study made by Paul and 
Cooksey (1979, 1981a,b) of an unidentified marine Chlamydomonas spe-
cies. This species has an active periplasmic asparaginase, which deami-
dates exogenous asparagine at the cell surface. The enzyme is induced 
by nitrogen deprivation and parallels glutamine synthetase in its regula-
tion, being repressed by high levels of N H 4
+
, N 0 3 ~ , or asparagine (Paul, 
1982). High levels of asparagine and N H 4
+
, but not of N 0 3 ~ , induce 
synthesis of glutamatedehydrogenase. 
Urea and Uric Acid Metabolism 
Urea can be used as the sole nitrogen source by C. reinhardtii (Sager and 
Granick, 1953) and appears to be taken up by an active transport mecha-
nism (Williams and Hodson, 1977). Dagestad et al. (1981) have sug-
gested that exogenous urea is actively transported into the chloroplast, 
where it forms a large nonmetabolic pool, and that urea catabolism 
occurs from a separate metabolically active pool. Urea is converted to 
N H 3 in a two-step process mediated by an enzyme complex designated 
in its entirety as A T P : urea amidolyase (EC 6.3.4.6) and consisting of 
two separable activities, urea carboxylase and allophanate hydrolyase 
(Thompson and Muenster, 1971; Whitney and Cooper, 1973; Hodson et 
al., 1975). The first reaction involves condensation with H C 0 3 " to form 
allophanate (Figure 6.3), which is then hydrolyzed to release H C 0 3 ~ and 
N H 3 . Overall, the reaction is the same as urea hydrolysis by urease (EC 
3.5.1.5), but the amidolyase enzyme differs from urease in requiring 
A T P , M g
2 +
, and K
+
 and in being sensitive to avidin (Leftley and Syrett, 
1973). Leftley and Syrett (1973) identified urea amidolyase in seven 
genera of Chlorophyceae, including C. reinhardtii, but found urease 
instead in representatives of four other algal classes. Both urea carbox-
ylase and allophanate lyase activities can be induced by urea or acet-
amide in NH 4
+
-dep r ived cultures and are repressed by N H 4
+
 (Hodson et 
al., 1975; Semler et al., 1975). Induction can occur at any stage in the cell 
cycle on addition of urea or acetamide and does not seem to have an 
obligatory temporal link to gametogenesis, which is also induced by 
ammonium withdrawal (see Chapter 4) . Xanthine is taken up by an 
active transport system and is assimilated by means of xanthine dehy-
drogenase, whose activity is induced by xanthine and other purines and 
is repressed by ammonium (Fernandez and Cârdenas, 1981a; Pérez-
Vicente et al., 1988). 
2 4 4 6. Metabolism 
Uric acid, or urate, can also be assimilated by C. reinhardtii cells. An 
active urate transport system is induced by transfer to urate-containing 
medium and is inhibited by N H 3 , darkness, and metabolic inhibitors 
(Pineda and Cârdenas, 1985). The first step in urate assimilation is medi-
ated by urate oxidase, which has been characterized by Pineda et al. 
( 1984a,b). Their results are consistent with aerobic purine catabolism 
following the pathway common to plants, animals, and many microor-
ganisms, which involves adenine, hypoxanthine, xanthine, urate, allan-
t o i c and allantoate, all of which can be used as nitrogen sources by C. 
reinhardtii. 
Urea, acetamide, and thiourea all induce synthesis of acetamidase, 
which catalyzes the hydrolysis in vitro of many different amides to acid 
plus ammonia (Gresshoff, 1981a,b). Acetamidase-deficient mutants fail 
to grow on formamide, acetamide, propionamide, or butyramide, sug-
gesting that this enzyme is a general amidase in C. reinhardtii cells 
(Hodson and Gresshoff, 1979). Ammonium represses synthesis of 
acetamidase (Gresshoff, 1981b). 
Arginine Biosynthesis 
The arginine biosynthetic pathway was one of the earliest metabolic 
processes to be studied in Chlamydomonas (Hudock, 1962, 1963; Eber-
sold, 1962), and was found to resemble the pathway in yeast and Neu-
rospora (Figure 6.4). Auxotrophic mutants are now known for six of the 
eight enzymes in this pathway (for complete references see Chapter 11). 
The mutants arg-1, arg-9, and arg-10 can grow if supplied with arginine, 
ornithine, or citrulline, while arg-7 and arg-8 have an absolute require-
ment for arginine. A s expected from their block at the ornithine carba-
moyltransferase step, the two allelic mutants at the arg-4 locus are un-
able to grow on ornithine, but they are also unable to use citrulline 
(Loppes , 1969a; Loppes and Heindricks, 1986). The reason for this is 
unclear. 
Acetylglutamate kinase is the allosteric enzyme of the pathway and is 
inhibited by arginine through a feedback loop (Farago and Denes, 1967). 
Ornithine carbamoyltransferase is constitutively expressed (Strijkert 
and Sussenbach, 1969). According to Hudock (1963), argininosuccinate 
lyase is strongly repressed by arginine, but Sussenbach and Strijkert 
(1969a) did not observe any relation between the activity of the lyase and 
the arginine concentration in the cell. Sussenbach and Strijkert also 
reported that high exogenous concentrations of ornithine inhibited 
growth of wild-type or arg-1 cells and led to the intracellular accumula-
tion of argininosuccinate. They postulated that arginyl-tRNA is the ac-
tual corepressor of argininosuccinate lyase, with argininosuccinate hav-
ing a compensatory inhibitory effect on arginyl-tRNA synthetase. Thus 
high concentrations of arginine inhibit step Β of the pathway shown in 
Figure 6.4, lowering production of the various precursors, including 
argininosuccinate. This causes decreased inhibition of arginyl-tRNA 
Nitrogen Assimilation 245 
glutamate α-ketoglutarate 
acetylglutamic semialdehyde 
C arg-1 
D arg-9 
glutamate acetylornithine 
γ 
acetylglutamyl phosphate 
A 
acetylglutamate ornithine 
F arg-4 
arginine 
H arg-Z 
argininosuccinate 
G arg-8 
• citrulline 
Enzymes: 
A: acetylglutamate synthetase (amino acid acetyltransferase) 
B: acetylglutamate kinase 
C: acetylglutamyl phosphate reductase 
D: acetylornithine aminotransferase 
E': acetylornithine glutamate transacetylase (glutamate acetyltransferase) 
F: ornithine carbamoyltransferase 
G: argininosuccinate synthetase 
H: argininosuccinate lyase 
Figure 6.4 Pathway of arginine biosynthesis, showing arginine-requiring mutants of C. 
reinhardtii. Adapted from Loppes and Heindricks (1986). 
synthetase, producing more arginyl-tRNA, which in turn represses syn-
thesis of the lyase. A s arginine is utilized in growth, eventually feedback 
inhibition of the early pathway is relieved, argininosuccinate accumu-
lates, arginyl-tRNA synthetase is inhibited, and the lyase is derepressed, 
leading to arginine production. Nevertheless, this model should be con-
sidered with caution since it is based partly on the assumption that arg-2 
lacks argininosuccinate synthetase (Hudock, 1963) rather than arginino-
succinate lyase (Strijkert et al., 1973). 
Approximately half the arginine-requiring mutants recovered from 
M N N G (iV-methyl-iV'-nitro-N-nitrosoguanidine) mutagenesis have 
proved to be arg-7 alleles and defective in argininosuccinate lyase activ-
ity (Strijkert et al., 1973; Matagne, 1976; Matagne and Vincenzotto, 
1979). This enzyme appears to be a homomultimer, composed of several 
(probably four) identical subunits of approximately 50 kDa (Farrell and 
Overton, 1987). A presumptive 39-kDa subunit (Matagne and Schlosser, 
1977) now appears to be an unrelated protein. Antibody to the 50-kDa 
subunit inhibits argininosuccinate lyase, whereas antibody to the 39-kDa 
246 6. Metabolism 
protein does not (Farrell and Overton, 1987). Loppes et al. (1972; see 
also Loppes and Matagne, 1972) showed that certain combinations of 
the arg-7 mutants are capable of intragenic complementation. The en-
zyme formed in these diploids is typically less active than its wild-type 
counterpart, but it is adequate to support growth of the diploids on 
minimal medium with no exogenous arginine. It is also more heat-labile 
than the wild-type enzyme. Matagne (1976, 1977) also investigated 
argininosuccinate lyase in diploids formed from arg-7 pab-2
+
 x arg-7
+ 
pab-2 crosses, in which some of the enzyme formed is presumed to 
contain both mutant and wild-type subunit s. Again, reduced activity and 
heat sensitivity were observed at least with some alleles. Similar results 
were obtained in triploid crosses among arg-7 and wild-type diploid and 
haploid cells in various combinations (Matagne andVincenzotto, 1979). 
T w o alleles, arg-7-1 and arg-7-6, which are unable to complement any 
other allele in diploid combinations, were shown to make an arg-7 gene 
product able to interfere positively or negatively with other mutant prod-
ucts in triploids (Matagne and Vincenzotto, 1979; Matagne, personal 
communication). Further analysis with additional alleles resulted in a 
composite recombination-complementation map for the arg-7 locus 
(Matagne, 1978) which is reproduced in the section on arg-7 in Chap-
ter 11. 
Exogenous arginine is actively taken up by wild-type Chlamydomo-
nas cells. Kirk and Kirk (1978a,b) confirmed that this uptake was in-
volved a specific transport system and that arginine was the only amino 
acid with carrier-mediated uptake. Arginine uptake is repressed strongly 
by N H 4
+
 (Loppes and Strijkert, 1972) and to a lesser extent by nitrate, 
urea, and leucine (Kirk and Kirk, 1978b). These results offer an explana-
tion for Loppes 's (1969a) observation that arginine auxotrophs were 
more easily isolated on medium low in N H 4
+
. Having isolated five new 
arg-7 alleles that were indeed sensitive to N H 4
+
, Loppes (1970b) ques-
tioned why the original arg-7 mutant isolated by Gillham (1965a) was 
insensitive. His conclusion was that this was a double mutant, carrying 
both arg-7 and a determinant producing insensitivity to N H 4
+
. Genetic 
analysis suggested that the two factors were closely linked (2.4 recombi-
nation units), and that a similar two-gene alteration could account for 
lack of N H 4
+
 sensitivity in the other original mutants, arg-1, arg-2y and 
arg-4. 
Sulfur Metabolism 
Exogenous sulfate is assimilated (Figure 6.5; see also Siegel, 1975) by 
reaction with A T P to form 3'-phosphoadenosine 5'-phosphosulfate 
( P A P S ) , in a two-step process catalyzed by ATP-sulfurylase ( A T P : 
sulfate adenylyltransferase) and APS-kinase ( A T P : adenylylsulfate 3'-
phosphotransferase). The latter enzyme has been characterized from C. 
reinhardtii by Jender and Schwenn (1984; see also Schwenn and Jender, 
Vitamins and Cofactors 247 
sulfate esters 
C3 + C4 
S 0 4
=
— • APS — • P A P S — • [ -S-S03
_
] —»-[-S-S"] -X amino acids 
[1] [2] [3] [4] 
- so 3= 
Enzymes: 
1 : ATP sulfurylase 
2: APS kinase 
3: APS:thiol sulfotransferase 
4: thiosulfonate reductase 
Figure 6.5 Pa thway of sulfate assimilat ion. 
1981). Schwenn and Schriek (1984) reported that isolated APS-kinase 
from Chlamydomonas was stimulated by spinach thioredoxin f, but they 
obtained no evidence that this compound was involved in reduction of 
P A P S to sulfite as occurs in bacteria. Instead, thiosulfate appears to be 
the primary reduction product in Chlamydomonas and other algae (Hod-
son and Schiff, 1971). The reduction step can proceed in vitro with 
reduced pyridine nucleotides or sulfhydryl reagents as reductants and 
requires two separable enzyme fractions, APS: th io l sulfotransferase 
and thiosulfonate reductase. Chlorella mutants deficient in these activi-
ties have been identified (Hodson et al., 1971), but comparable mutants 
have not yet been isolated in Chlamydomonas, nor have extensive in-
vestigations of biosynthesis of sulfur-containing amino acids or other 
compounds been made. Sulfolipid formation can occur with either P A P S 
or sulfite as sulfonyl donor in a cell-free system from Chlamydomonas, 
but the pathway utilized in vivo is not yet fully elucidated (Hoppe and 
Schwenn, 1981). Isolation of an APS-sulfohydrolase fraction capable of 
hydrolyzing A P S to nucleotides, including adenosine-5'-phosphorami-
date ( A P N ) , 5 -AMP, adenosine, and c A M P , was reported by Kuhlhorn 
and Schmidt (1980). 
Certain organic compounds can presumably also be used as sulfur 
sources by sulfate-deprived Chlamydomonas, but the range of suitable 
substrates has received relatively little attention. Lien and Schreiner 
(1975) characterized an arylsulfatase from C. reinhardtii that was dere-
pressed by sulfate starvation. Like the phosphatases and carbonic anhy-
drase, this enzyme appears to be localized close to the cell surface. 
Vitamins and Cofactors 
Despite the utility of vitamin-requiring auxotrophic mutations for ge-
netic analysis, very little is known specifically about Chlamydomonas 
248 6. Metabolism 
regarding the synthesis of vitamins and other cofactors. Wild-type C. 
reinhardtii strains appear to require no supplements in the medium. 
However , many algae isolated in nature require vitamin B j 2 and/or thi-
amine (see Provasoli and Carlucci, 1974), and within the genus Chlamy-
domonas several natural auxotrophs for vitamin B | 2 are known. These 
include C. chlamydogama (Bold, 1949a; Trainor, 1958), C. hedleyi (J. J. 
L e e et al., 1974), C. pallens (Pringsheim, 1962), and C. Pulsatilla 
(Droop, 1961). The existence of auxotrophic species suggests that isolat-
ing comparable mutants in C. reinhardtii might be possible, although so 
far none has been obtained. 
Thiamine-requiring auxotrophic mutants are known both in C. 
reinhardtii and C. eugametos; they have been useful genetic markers but 
have not been investigated extensively in a biochemical sense (see Chap-
ter 11). Presumably thiamine biosynthesis follows the pathways known 
in higher plants (see Ebersold, 1962). Some mutants, such as C. 
reinhardtii thi-2, can grow on vitamin thiazole (4-methyl-5-/3-hydroxy-
ethyl thiazole) plus the pyrimidine moiety of thiamine (2-methyI-4-
amino-5-ethoxy-methyl pyrimidine), while others, such as thi-l, require 
intact thiamine. The thi-3 and thi-4 mutants can use either thiamine or 
thiazole and do not require pyrimidine, but they differ in their sensitivity 
to the analogs oxythiamine and pyrithiamine, thi-3 being sensitive and 
thi-4 being insensitive to these compounds in combination. Mutants re-
sistant to pyrithiamine have also been isolated both in C. eugametos and 
C. reinhardtii. McBride and Gowans (1967) showed that pyrithiamine-
resistant mutants of C. eugametos were impaired in thiamine uptake, 
and that the combination of the resistance mutation with a thiamine-
requiring auxotrophy mutation was lethal. The single pyrithiamine resis-
tance locus identified in C. reinhardtii maps very close to thi-4, an auxo-
trophic mutation, but has not been investigated physiologically (Smyth 
et al., 1975). 
Eversole (1956) isolated nicotinamide auxotrophs in C. reinhardtii and 
showed that some but not all of these were able to use 3-hydroxyan-
thranilic acid, quinolinic acid, or kynurenine to replace nicotinamide 
(see Chapter 11 for details). Gowans (1960) first reported isolation of 
nicotinamide auxotrophs in C. eugametos. Nakamura and Gowans 
(1964) isolated a mutant resistant to the analog 3-acetyI pyridine and 
showed that this mutant excreted excess nicotinic acid into the medium, 
suggesting that nicotinic acid synthesis in the mutant might be dere-
pressed. In a second paper (1965), they reported a nonallelic mutation 
conferring partial resistance to 3-acetyl pyridine and producing an inter-
mediate level of nicotinic acid excretion. Neither resistance mutation 
appeared to be allelic with any of the nicotinamide auxotrophy muta-
tions. The auxotrophic mutations in C. eugametos were grouped into 
five independent loci (Nakamura and Gowans, 1965). None could use 
hydroxyanthranilic acid, kynurenine, or other precursors in the tryp-
Physiological Ecology 249 
tophan-nicotinic acid pathway. Nicotinic acid at high concentration 
supported growth of all five, and quinolinic acid was effective for two 
groups of mutants. A secondary mutation, mod-1, which inhibits nico-
tinic and quinolinic acid utilization by nic-5 and nic-6 mutants, probably 
affects uptake of these compounds rather than their metabolism (Naka-
mura and Gowans, 1965, 1967). Excess K C l or other salts partially alle-
viated the mod-1 effect. In the last paper from Gowansand co-workers 
on nicotinamide metabolism, Uhlik and Gowans (1974) confirmed that 
synthesis of nicotinic acid in C. eugametos begins with tryptophan, as in 
mammals and ascomycetes, rather than with glyceraldehyde-3-phos-
phate and aspartic acid as in higher plants. 
Mutants requiring p-aminobenzoic acid are also known in both C. 
eugametos/C. moewusii and C. reinhardtii but have received very little 
study beyond their use as genetic markers (Chapter 11). T w o mutants of 
C. eugametos that responded to exogenous pyridoxine and two respond-
ing to folic acid were reported by Wethereil and Krauss (1957) but were 
incompletely characterized and apparently were not preserved. The vi-
tamin requirements did not appear to be specific, in any case. 
Physiological Ecology 
Most metabolic studies with Chlamydomonas have focused on behavior 
under laboratory conditions, often far removed from the situation to be 
found in nature, while more ecological investigations have tended to 
concentrate on population dynamics (e .g . , Archibald and Bold, 1976; 
Boyd, 1972; Cunningham and Maas, 1978; Cunningham and Nisbet, 
1980; Happey-Wood, 1980; Merrett and Armitage, 1982; Richards and 
Happey-Wood, 1979) or response to specific environmental agents (Ta-
ble 6.7). A few studies have at least begun to set the physiological 
characteristics of the organism into a natural context and to relate obser-
vations on nitrogen metabolism, photosynthesis, carbon balance, etc. 
Especially noteworthy among the older studies are the papers on game-
togenesis and enzyme synthesis in synchronous cultures by Kates and 
Jones (1964a, 1966, 1967). Hudock et al. (1971) studied phosphate, argi-
nine, and acetate limitation in cells grown in a chemostat culture. Kroes 
(1971-1973) investigated excretion of extracellular products and growth 
interactions between Chlamydomonas globosa and Chlorococcum ellip-
soïde urn under different environmental conditions. Cohen and Parnas 
(1976) treated carbon metabolism, specifically synthesis of storage mate-
rials, under natural (diurnal) conditions in a theoretical model. Bollman 
and Robinson (1977) measured the ability of C. segnis and other algae to 
assimilate organic acids and compared these rates with the potential 
assimilation by bacterial populations in natural waters. Slawyk et al. 
(1977) measured carbon and nitrogen turnover rates in marine Chlamy-
250 6. Metabolism 
T a b l e 6.7 Stud ies of Effects of Pollution 
on Chlamydomonas S p e c i e s
3 
Pollutant Species studied Reference 
Arsenic 
Cadmium 
Lead 
Heavy metals 
Mercury 
Sulfur dioxide 
Nitrogen dioxide 
Chlorine 
Sulfite 
Naphthalene 
Crude oil extracts 
Polychlorinated 
biphenyls (PCBs) 
Organophosphates 
and other 
insecticides 
C. reinhardtii 
Freshwater species 
C. reinhardtii 
C. reinhardtii 
Freshwater species 
C. variabilis 
C. sp. (Carolina Biol.) 
C. reinhardtii 
C. reinhardtii 
C. reinhardtii 
Marine species 
C. reinhardtii 
C. angulosa 
C. Pulsatilla 
C. angulosa 
C. reinhardtii 
C. reinhardtii 
Freshwater species 
C. reinhardtii 
Planas and Healey (1978) 
Christensen and Zielski (1980) 
Fennikoh et al. (1978) 
Ahlf et al. (1980); Irmer et al. (1986) 
Foster (1982) 
Delcourt and Mestre (1978) 
Knowles and Zingmark (1978) 
Ben-Bassat et al. (1972) 
Wodzinski and Alexander (1978) 
Wodzinski and Alexander (1980) 
Hirayama and Hirano (1980) 
Stamm (1980) 
Soto et al. (1975a, 1975b, 1979b); 
Hellebust et al. (1982, 1985); 
Hutchinson et al. (1981, 1985) 
Hsiao (1978) 
Soto et al. (1975a, 1977, 1979a); 
Hellebust et al. (1982, 1985); 
Hutchinson et al. (1981, 1985) 
Vandermeulen and Lee (1986) 
Gresshoff et al. (1977); 
Mahanty and Gresshoff (1978); 
Conner (1981) 
Christensen and Zielski (1980) 
Birmingham and Colman (1977); 
Netrawali et al. (1986) 
" Also see Palmer (1969) for early references. 
domonas, and Barrett and Koch (1982) studied nitrogen utilization in 
Chlamydomonas and other green algae from rice fields. Derivation of 
nitrogen and other nutrients from symbiotic or commensal relationships 
is discussed by Goff and Stein (1978) for Chlamydomonas associated 
with jelly surrounding salamander eggs, by Saks (1982) for C. provasolii, 
an endosymbiont of a foraminifer, and by Gyurjân et al. ( 1984a,b), who 
created an artificial association of Chlamydomonas with a nonsymbiotic 
nitrogen-fixing bacterium, Azotobacter. Phosphorous assimilation in 
natural populations of Chlamydomonas and other algae has been studied 
by Currie and Kalff (1984). Harris and Piccinin (1983) have dealt with 
overall photosynthetic rates and with phosphorous and carbon metabo-
lism in C. reinhardtii, and Necas and Tetik (1985) have described effects 
of nitrogen limitation in C. geitleri. Gleason and Baxa (1986) have stud-
ied the effects of the natural algicide cyanobacterin, produced by the 
cyanobacterium Scytonema hofmanni, on various algae including C. 
reinhardtii. 
Excretion of Metabolic Products into the Medium 251 
Ion Transport 
Surprisingly little is known either about the nutritional requirements of 
Chlamydomonas for specific ions, or about their entry into the cell. 
Eppley (1962) and MacRobbie (1974) reviewed ion transport in algae but 
neither mentioned any studies specifically on Chlamydomonas. Moss et 
al. (1971) reported that C. reinhardtii could grow with strontium replac-
ing calcium in the culture medium. Mutants resistant to cadmium, cop-
per, and zinc and mutants resistant to cobalt and nickel have been iso-
lated by Collard (1985). Tolerance to high levels of chloride salts (NaCl , 
K C l , L i C l ) is increased by exposure of C. reinhardtii cells to taurine 
(Reynoso and Gamboa, 1982; Gamboa et al., 1985), but the mechanism 
of this effect remains to be fully elucidated. Growth of C. reinhardtii in 
proline also induces salt tolerance (Reynoso-Granados et al., 1985). 
Some halotolerant Chlamydomonas species, such as C. Pulsatilla, accu-
mulate intracellular glycerol in response to osmotic stress (Hellebust, 
1985b; see also Ben-Amotz and Avron , 1983). Chlamydomonas 
reinhardtii also can accumulate and excrete glycerol when grown on 
>100 mM K C l or NaCl or 200 m M sucrose ( H . D . Husic, personal 
communication). Chlamydomonas Pulsatilla has also been used in stud-
ies of sodium effects on uptake of other nutrients (Hellebust, 1985a). 
This species grows over a moderately wide range of salinity but does not 
require any minimal concentration of sodium for growth, in contrast to 
Dunaliella tertiolecta, which has an absolute requirement for sodium for 
uptake of phosphate and methylamine. 
In an early study, Teichler-Zallen (1969) analyzed the effects of man-
ganese on chloroplast structure and photosynthetic function. Sunda and 
Huntsman (1985) have investigated manganese uptake in a marine 
Chlamydomonas species. Zinc uptake in C. variabilis has been studied 
by Bates et al. (1982, 1985) and by Harrison et al. (1986). Bates et al. 
proposed that zinc is bound by cellular polyphosphate and accumulates 
in young cultures; as phosphate decreases in older cultures, zinc is 
released and subsequently interferes with cell division. Polley and Doc-
tor (1985) have isolated C. reinhardtii mutants that require high levels of 
potassium for growth and have shown that these mutants are specifically 
defective in transport activity. These investigations should be the basis 
for continued exploitation of the potential of Chlamydomonas for ge-
netic analysis of transport mechanisms. 
Excretion of Metabolic Products into the Medium 
A s mentioned earlier, periplasmic enzymes such as phosphatases, sulfa-
tases, and carbonic anhydrase are excreted into the culture medium by 
wild-type cells, and they reach even higher concentrations in cultures of 
cell wall-deficient mutants. Excretion of various other compounds has 
252 6. Metabolism 
been reported from several species (see Fogg, 1962, forreview), but a 
systematic investigation of the physiological aspects of these processes 
has not been made. An early study by Allen (1956) documented excre-
tion of glycolate, oxalate, and pyruvate by C. reinhardtii, C. eugametos, 
C. moewusii, a species identified as C. pseudogloea (C. pseudagloë?), 
and two unidentified species isolated from sewage oxidation ponds. Col-
lins and Kalnins (1967) reported excretion of several α-keto acids by C. 
reinhardtii. Kroes (1972a,b), using C. globosa, reported finding steam-
volatile acids, water-soluble yellow phenolic compounds, unidentified 
lipophilic substances, proteins, and polysaccharides in the medium after 
culture. Excretion of vitamins, especially folic acid, biotin, and pan-
tothenic acid, was reported by Aaronson et al. (1977). Excretion of 
c A M P , up to 85% of the total synthesized by wild-type C. reinhardtii 
cells, was reported by Bressan et al. (1980). Dissolved amino acids and 
sugars were found in culture media after growth of C. reinhardtii by 
Vogel et al. (1978). Brown and Geen (1974) reported increased excretion 
of ethanol-soluble organic acids from cells grown in green or white light 
and more protein and amino acids excreted from cells grown in blue 
light. Cristofalo et al. (1962) noted that culture media of C. moewusii 
took on a yellow color, which they tentatively identified as a mixture of 
organic acids, probably the result of natural excretion rather than de-
composition. 
Why So Few Auxotrophs? 
The range of auxotrophic mutations identified in Chlamydomonas is 
severely limited: the only amino acid auxotrophs are C. reinhardtii mu-
tants requiring arginine, and only a few auxotrophs for vitamin cofactors 
have been identified (nicotinamide, thiamine, p-aminobenzoic acid). 
Purine-requiring mutants have been reported in C. eugametos (Gowans, 
1960) but not in C. reinhardtii. Several possible explanations have been 
advanced for the paucity of auxotrophs, including fundamental differ-
ences in metabolic pathways of green plants compared to those of fungi 
and bacteria, differences in inducibility or repression of biosynthetic 
enzymes, permeability barriers, and extensive gene duplication. 
Li et al. (1967) surveyed the spectra of auxotrophic mutations in a 
variety of organisms and concluded that Chlamydomonas, a moss (Phys-
comitrella), and a higher plant (Arabidopsis) all were similarly poor in 
auxotrophs compared to bacteria and fungi. N o differences in mutagene-
sis methods or selection techniques could be found that would account 
for this discrepancy. Kirk and Kirk (1978a,b) were unable to detect 
active (carrier-mediated) uptake for any amino acid except arginine in C. 
reinhardtii, although Loppes (1969a) had reported that leucine at least 
was taken up sufficiently well to serve as a sole nitrogen source. The rate 
of uptake of leucine observed by Kirk and Kirk could be accounted for 
by passive diffusion but was indeed marginally adequate to support 
Effects of Herbicides and Metabolic Inhibitors 253 
growth (a 1 m M solution was sufficient to provide 10 nmole/min per mg 
of cellular protein). The nonmetabolizable amino acid analog 2-amino-
isobutyric acid appears to be actively transported, at least to some ex-
tent (Hasnain and Upadhyaya, 1982). Proline utilization in response to 
high salt concentration was reported by Reynoso and Gamboa (1982). 
Glutamine can be used as the sole nitrogen source by C. reinhardtii (see 
Table 6.5), and growth of C. reinhardtii is inhibited by glyphosate, which 
blocks aromatic amino acid biosynthesis (Gresshoff, 1979), and by chlor-
sulfuron, which interferes with isoleucine-valine synthesis (Hartnett et 
al., 1987). Thus there is substantial evidence in favor of uptake of a 
number of amino acids. Kirk and Kirk (1978b) suggested that previous 
searches for auxotrophs failed because amino acids were not provided at 
sufficiently high concentration in the nonselective medium to which cells 
were exposed following mutagenesis. In the 10 years following publica-
tion of this paper, no one has come up with additional amino acid auxo-
trophs in C. reinhardtii', the Kirks, whose primary research interest is 
Volvox, have not pursued this project ( D . Kirk, personal communica-
tion). 
Nakamura et al. (1981) showed that methionine could competitively 
relieve inhibition by methionine sulfoximine in C. reinhardtii, suggesting 
that at least some methionine was being taken up. However , they too 
were unable to isolate an auxotrophic mutant and found in fact that cells 
on methionine medium were killed by daylight fluorescent lights. They 
therefore postulated a photodynamically produced toxicity that might 
inhibit recovery of mutants on medium high in methionine and, by exten-
sion, perhaps other compounds as well. A mutant that was hypersensi-
tive to this methionine-mediated light damage was isolated by Nakamura 
et al. (1979, 1981; Nakamura and Lepard, 1983). Catalase and superox-
ide dismutase suppressed the methionine photoinactivation, leading Ta-
kahama et al. (1985) to conclude that the mechanism of damage is forma-
tion of peroxide and O 2 " . Photodynamic toxicity was also proposed as a 
mechanism for tryptophan sensitivity of some C. eugametos and C. 
reinhardtii strains (Nakamura et al., 1979). 
In a later publication, Nakamura et al. (1985) reported their failure to 
isolate arginine auxotrophs in C. eugametos. This species is capable of 
at least limited arginine uptake, since arginine alleviates inhibition by 
canavanine, but assimilation of radioactive arginine was considerably 
lower than in C. reinhardtii. Thus at least in this case, the failure to 
isolate auxotrophs probably does result from inefficient uptake of exoge-
nous amino acid. 
Effects of Herbicides and Metabolic Inhibitors on Chlamydomonas 
The use of Chlamydomonas as a model system for understanding the 
metabolism of higher plants, and particularly its potential as a vehicle for 
genetic engineering, requires that its response to herbicides and other 
254 6. Metabolism 
inhibitors be known. Table 6.8 summarizes the literature on effects of 
various types of herbicides on Chlamydomonas species, chiefly C. 
reinhardtii. Cain and Cain (1983) have studied the sensitivity of vegeta-
tive cells and germinating zygospores to a variety of herbicides. Table 
6.9, which is reproduced from Mottley and Griffiths (1977), deals mainly 
with inhibitors of photosynthesis and respiration, some of which may 
also be useful as herbicides. Table 6.10, expanded from McBride and 
Gowans (1970), compares effects of analogs of amino acids, vitamins, 
and other metabolic compounds on C. eugametos, C. moewusii, and C. 
reinhardtii. Inhibitors of nucleic acid and protein synthesis are discussed 
at greater length in Chapters 8 and 9, and additional information on 
preparation of test media containing inhibitors will be found in Chap-
ter 12. 
T a b l e 6.8 Herb ic ides T e s t e d on Chlamydomonas S p e c i e s 
Class of compounds Examples Action References 
Dinitrophenols, hydroxy- Dinoseb, Bromoxynil Inhibit respiration and Cullimore (1975); Hess (1980); 
benzonitriles, penta- photosynthesis Fedtke (1982) 
chlorophenol 
Ureas, uracils, triazines, DCMU, Bromacil, Atra- Block photosynthetic Loeppky and Tweedy (1969); 
triazinones zine, Metribuzin electron transport Lien et al. (1977); Senger 
(1977); Oettmeier et al. 
(1981); Fedtke (1982); 
Shochat et al. (1982); 
Galloway and Mets (1982, 
1984); Pucheu et al. (1984); 
Maule and Wright (1984); 
Erickson et al. (1984a, 
1985a) 
Bipyridinium derivatives Diquat, Paraquat Cause lethal 0 2 free radical 
formation in photosyn-
thesis 
Cullimore (1975); Hess (1980); 
Fedtke (1982) 
Nitrodiphenylethers Nitrofen, Bifenox Cause free radical formation 
in photosynthesis 
Hess (1980); Fedtke (1982); 
Ensminger and Hess 
(1985a,b); Ensminger et al. 
(1985)Pyridazinones, amino- Fluridone, Amitrole Block carotenoid synthesis Vance and Smith (1969); Hess 
triazole, other "bleach- (1980); Fedtke (1982) 
ing" herbicides 
N-Phenylcarbamates Chlorpropham Inhibit microtubule assem-
bly 
Cullimore (1975); Hess (1980); 
Fedtke (1982); Maule and 
Wright (1983, 1984) 
Dinitroanilines Oryzalin, Trifluralin Inhibit microtubule assem-
bly 
Hess (1979, 1980); Quader 
and Filner (1980); Fedtke 
(1982); Strachan and Hess 
(1983); James et al. (1987) 
Phosphoric amides Amiprophos methyl Prevent tubulin synthesis Collis and Weeks (1978); 
Quader and Filner (1980); 
Fedtke (1982) 
N-Phosphonomethylglycine Glyphosate Blocks aromatic amino acid 
biosynthesis 
Gresshoff (1979); Hess (1980); 
Maule and Wright (1984) 
Chloroacetanilides Alachlor Metabolic inhibition, site 
unknown 
Hess (1980); Fedtke (1982) 
Thiocarbamates Butylate, EPTC, Molinate Block fatty acid biosynthe-
sis 
Hess (1980) 
Effects of Herbicides and Metabolic Inhibitors 255 
T a b l e 6.8 {continued) 
Class of compounds Examples Action References 
Miscellaneous other com-
pounds 
M C P A , MCPB 
Bensulide 
Dichlobenil 
Diphenamid 
Aerotex 3470 
Respiratory inhibitors 
Inhibits polysaccharide 
synthesis 
Kirkwood and Fletcher 
(1970); Maule and Wright 
(1984) 
Hess (1980) 
Cullimore (1975); Hess (1980) 
Loeppky and Tweedy (1969) 
Moody et al. (1981) 
T a b l e 6.9 Toxic L e v e l s of Var ious Inhibitors for C. reinhardtii
ab 
Minimum inhibitory concentration for a given condition 
Compound Phototrophic Mixotrophic Heterotrophic 
Energy transfer inhibitors 
Trimethyltin chloride 20.5 10.2 10.2 
Triethyltin sulfate 3.9 3.5 3.0 
Tri-rt-propyltin chloride 1.8 1.8 1.1 
Tri-rt-butyltin chloride 1.2 1.2 0.3 
Triphenyltin chloride 1.0 0.8 0.1 
Tri-H-butyltin oxide 0.2-1.7 NT <0.2 
Tricyclohexyltin hydroxide 2.6 2.6-26.0 0.3-2.6 
Octylguanidine 19.3 16.9 14.5 
Dodedylguanidine 7.6 7.6 7.6 
Galegine sulfate >113 113-284 425 
Oligomycin >25 /ig/ml >25 /Ltg/ml 10 /Ag/ml 
Phlorizin >229 >229 >229 
Venturicidin >126 >126 32-126 
DCCD 121-242 >242 >242 
Dio-9 25-50 /ig/ml 25-50 /Ltg/ml 1-5 μg/ml 
Robenzidine >75 >75 3-30 
Aurovertin >10^g/ml >10 /Ltg/ml 1-5 /xg/ml 
Uncouplers 
DNP >1358 >1358 >1358 
Atebrin >106 >106 >106 
CCCP 14.6 14.6 8.5 
1799 90 >128 51 
Sodium arsenate >1603 >1603 >1603 
Tetraphenylboron >292 >292 >292 
TTFB 32-79 32-79 32-79 
Adenine nucleotide translocase inhibitors 
Atractyloside >119 >119 >119 
Rhodamine 6G 111-222 22-111 <22 
Ionophores 
Valinomycin >45 >27 9 
Nigericin 5 μ-g/ml 7.5 jug/ml 5 /*g/ml 
Gramicidin >50 /Lig/ml >50 /xg/ml >50 /xg/ml 
Dicyclohexyl-18-crown-6 >269 >269 54 
(continued) 
256 6. Metabolism 
Minimum inhibitory concentration for a given condition 
Compound Phototrophic Mixotrophic Heterotrophic 
Electron transport inhibitors 
Potassium cyanide >3840 >3840 >3840 
Antimycin A >18 >18 0.09-0.9 
Amytal >H05 > 1105 >1105 
Rotenone >127 >127 >127 
D N A and protein synthesis inhibitors 
Proflavin >3l4 >314 >314 
Acriflavin >100 μg/m 1 >100μg/ml 10 μg/ml 
Acridine orange >419 >419 140 
Spectinomycin 45 60 60 
Rifampicin >358 >358 287 
Ethidium bromide 19.0 19.0 12.7 
Detergents 
Sodium dodecyl sulfate 173-347 347-694 173-347 
Sodium deoxycholate 1449 1449 1208 
Cetyltrimethylammonium bromide 7-14 14-27 <7 
Miscellaneous 
Rhodamine Β 21-104 21-104 21-104 
Diethyltin dichloride >403 >403 >403 
Tetra-Aï-butyltin 29-144 3-29 3 
Di-A7-octyltin dichloride >240 120-240 24-120 
Di-H-butyltin diacetate 29-143 29-143 3-29 
Cl-methyl-di-A?-butyltin chloride 79 79 79 
" From Mottley and Griffiths (1977). For further information on triorganotins and /7-alkyl guanidines, see Mottley (1978). 
h
 All samples grown on solid media. Concentrations expressed in μΜ except where otherwise noted. N T , not tested. DCCD, 
Λ^ΛΓ-dicyclohexylcarbodiimide; DNP, 2,4-dinitrophenol; CCCP, carbonyl cyanide w-chlorophenylhydrazone; 1799, unidentified 
compound manufactured by DuPont; TTFB, 4,5,6,7-tetrachloro-2-trifluoromethylbenzimidazole. 
T a b l e 6.10 Sensit iv i ty of C. reinhardtii, C. eugametos, a n d C. moewus/7 to 
Metabo l ic A n a l o g s
3 b 
Concentration 
Compound tested C. eugametos C. moewusii C. reinhardtii 
3-Acetyl pyridine 0.8 mg/ml NT -
Allyl-D,L-glycine 1 mg/ml + + + 
2-Amino-3-phenylbutanoic acid 0.5 mg/ml - - + 
Aminopterin 0.5 mg/ml + + — 
L-3-Aminotyrosine HCl 0.5 mg/ml + + + 
8-Azaguanine 0.5 mg/ml + — 
4-Azaleucine 0.5 mg/ml + + + 
Benzimadazole 1 mg/ml - — 
Bromouracil 1 mM NT NT — 
Caffeine 20 mM NT — 
L-Canavanine sulfate 0.5 mg/ml + — 
2-Chloro-4-aminobenzoic acid 1 mg/ml + + + 
D,L-Ethionine 1 mg/ml + + 
T a b l e 6.9 (continued) 
Effects of Herbicides and Metabolic Inhibitors 257 
T a b l e 6.10 ( c o n t i n u e d ) 
Concentration 
Compound tested C. eugametos C. moewusii C. reinhardtii 
Fluoroacetate 10 mM NT NT -
Fluorouracil O.l mM NT NT -
D,L-p-Fluorophenylalanine l mg/ml + + + 
6-yV-Hydroxylamino purine l mg/ml + + + 
Imidazole 5 mM NT NT -
Indole 0.5 mg/ml - -
D,L-Methionine sulfoxide l mg/ml + - + 
L-Methionine-D,L-sulfoximine 0.5 mg/ml - + -
a-Methyl-D,L-methionine 0.5 mg/ml + + + 
D,L-Norleucine l mg/ml + + + 
D,L-Norvaline I mg/ml + + + 
Oxythiamine HCl 0.5 mg/ml + + + 
D,L-jS-Phenyllactic acid 1 mg/ml + + + 
Pyridine-3-sulfonic acid* - NT NT 
Pyrithiamine HBr 1 ug/ml - - -
D,L-Selenomethionine 0.25 mg/ml - - + 
D,L-Serine methylester HCl 0.5 mg/ml + + 
Sulfanilamide 1 mg/ml + + -
β-2-Thienylalanine 1 mg/ml + + 
" Modified from McBride and Gowans (1970), with additional information from Lawrence and Davies (1967), McBride and 
Gowans (1969), McMahon and Langstroth (1972), Flavin and Slaughter (1974), Hartfiel and Amrhein (1976), Wiseman et al. 
(1977a), Warr et al. (1978), and Gresshoff (1981b). 
h
 Test samples grown on agar under phototropic conditions. - , Sensitive; + , resistant at concentration tested; N T , not tested. 
' Concentration not specified in original paper; tests by the Chlamydomonas Genetics Center indicate that at least 0.5 mg/ml is 
required to kill wild-type C. eugametos.