ABSTRACT

A STUDY AND COMPARISON OF GLYCOLATE
METABOLISM IN TOBACCO AND GREEN ALGAE

by John L. Hess

The Glycolate Pathway in TobaccopLeaves:

Kinetic studies on the rate of glycolate biosynthesis
by tobacco leaves during 1“002 photosynthesis were run with
d-hydroxypyridinemethanesulfonate, an inhibitor of glycolate
oxidase. Although lL‘COZ fixation was reduced about 50 %.
the inhibitor did not affect the initial rate of 1“c incor-
poration into phosphoglycerate or glycolate, nor the Specific
activity of these products. The rate of glycolate formation
was slower than the rate of phosphoglycerate formation. At
time periods of 4 and 11 seconds, phOSphoglycerate was pre-
dominately carboxyl labeled and glycolate was uniformly
labeled. The specific activity of the carboxyl carbon of
phOSphoglycerate was about 100 fold higher than the Specific
activity of either carbon of glycolate, but after 60 seconds,
the’specific activities of the carbon atoms of both compounds
became nearly equal. The sulfonate blocked the glycolate

14C in glycolate rose from 3 % in the con-

pathway since the
trol to 50 z in the treated leaf, while the percent 1“0 in the
products of the glycolate pathway dropped from 32 % to 10 %.
These data emphasize the magnitude of the glycolate pathway

in higher plants, but do not indicate a second carboxylation

reaction in photosynthesis for glycolate production.

John L. Hess - 2

The Glycolate Pathway in Algae:

Enz es: No glycolate oxidase activity could be de-
tected in cell extracts from four different algae by mano-
metric. isot0pic, and Spectrophotometric assays. Isocitrate
dehydrogenase, NADH:glyoxylate reductase, and phospho-
glycolate phOSphatase were present. The absence of glycolate
oxidase is consistent with glycolate excretion by algae and
represents a major metabolic difference between higher plants
and algae.

Distribution of 1“0 Within Products Formed Duringlucog
Photosynthesis by Algae: For Chlorella and Chlamydomonas,
after 12 seconds photosynthesis, the carboxyl carbon contained

14

70 i to 80 % of the C in serine, but glycine was uniformly
labeled. Comparison of these results with similar experiments
with soybean leaves demonstrated that the complete glycolate
pathway does not function in algae.

The Effect of’d-hydroxymethanesulfonates on Photosyn-
thesis bz Algae: Addition of either 0.001 M‘NAhydroxy-
pyridinemethanesulfonate or’azhydroxymethanesulfonate to
Chlorella or Chlamydomonas produced four major alterations on
algal photosynthesis in 1“C02. At pH 8.3 the sulfonates re-
stored 002 fixation rates to those at pH 6.5. a three-fold

increase over the control. No change in rate occurred at

John L. Hess - 3

pH 6.5. The algae, in the presence of these inhibitors, did
not accumulate glycolate at either pH 6.5 or 8.3 as did the
tobacco. Both sulfonates caused a decrease in amino acid

14

synthesis, and a corresponding increase in the C accumula-
tion in the keto acids. Sugar phosphate metabolism was also
altered. since the<x-hydroxymethanesulfonates caused an in-
crease in these esters, particularly in ribulose-l,5-diphosphate.
It was concluded that these sulfonate inhibitors are not
specific for glycolate oxidase.

Growth and Photosynthesis of Algag in Red and Blue

 

gigggz Changes in absorption Spectrum and 1“(:02 fixation
patterns were observed with algae grown in red light C>600 mp)
or in blue light (“00-500 mu). When grown in blue light,
Chlorella and Chlamydomonas showed a consistent decrease in
their chlorOphyll a/b ratio. Chlamydomonas adapted to blue
light produced a larger amount of glycolate-140, while cells
adapted to red light labeled mainly sugar phosphates and
citric acid cycle products. At low light intensities (50-
100 ft-c), cells adapted to blue light accumulated 140 in
sugar phOSphates, while the cells grown in white formed mainly
products of dark fixation.

Manganese Deficient Chlorella: These cultures were

obtained after 20 days growth in a Mn+2 deficient medium. After

John L. Hess - #

lac in glycolate

photosynthesis for 10 minutes the percent
formed by normal algae was 30 %; but that formed by the defi-
cient algae was only 2 %. Simultaneously the percent Inc in
glycine and serine formed by the deficient algae increased.
These data suggest that serine is not formed from glycolate

in algae.

Glycolate Excretion and Uptake by_Algae: Uptake was
measured by the quantitative disappearance of glycolate in the
medium or by the appearance of 1“'0 labeled glycolate or
phOSphoglycolate in the cells. No time dependent uptake of
significant amounts of either glycolate or phosphoglycolate
was observed. Only 2 Z to 3 % of glycolate-Z-luc fed to
Chlorella and Scenedesmus was metabolized after 10 minutes in
the light. Glycolate excretion was maximal in 0.01 M bi-
carbonate and occurred in the light even in the presence of
10'5 M glycolate. Apparently algae were not capable of
glycolate metabolism as was the higher plant.

Comparison of Glycolate Metabolism in Algae and Tobacco‘
Leaves: It was concluded that there exists a basic difference
between the metabolism of glycolate by algae and by the higher
plant. Although in both plants, 3-phosphoglycerate was the
initial product of photosynthesis with the subsequent formation
of glycolate from the carbon reduction cycle, only in the higher
plant has glycolate metabolism via glycolate oxidase been found.

The algae. however, excrete their glycolate into the medium as

an apparent end-product.

A STUDY AND COMPARISON OF
GLYCOLIC ACID METABOLISM IN
TOBACCO AND GREEN ALGAE

By

John L. Hess

A THESIS

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY

Department of Biochemistry
1966

ACKNOWLEDGEMENTS

I especially thank Dr. N. E. Tolbert
whose creativity and requirements for ex-
cellence continue to affirm the current
excitement of modern biochemical research.
His encouragement, understanding and
guidance throughout the development of my
research program have been invaluable.

I am extremely thankful to Karen C.
Swanson for her excellent help. particularly
with the chromatography and radioauto-
graphy so essential to the work of this
thesis. I also thank Fang Hui Liao and P.
L. Ioungblood’ for their work with envir-
onmental factors and glycolate excretion.
and 0.1. Byard *for help with the glycolate
degradation technique.

I acknowledge the support and interest

of the National Science Foundation in this
research program.

*NSP Undergraduate Research Participant.
Summer. 196“.

*‘HBF Undergraduate Research Participant.
Summer 19 5.

ii

TABLE OF CONTENTS
INTRODUCTION 0 O 0 O 0 O O 0 C O 0 O 0 O O O 0 O 0

LITERATURE REVIEW . . . . .
Biosynthesis of Glycolate. . .
Biosynthesis of Glycine. . .
Biosynthesis of Serine . . .

The Glycolate Pathway in Higher Pla
The Glycolate Pathway in Algae . .
Glycolate Excretion and Uptake . .

nts .
Function of the Glycolate Pathway. . . .
t

0 O O 0 O 9 O O
O O O O O O O 0

Environmental Factors Affecting Glycola e
Formation. . . . . . . . . . . . . .

MATERIALS AND METHODS . . . . . . . . . . . . . .
Plant material 0 O O O O O O 0 O O O O C O 0

Chemicals. . . . .
Photosynthesis Experiments with Tobacco
Leaves . . . . . .

Photosynthesis Experiments with Algae. . .
Separation and Identification of Photo-
synthetic Products . . . .

Colorimetric Assays. . . . . . . . . . . . .
Degradation Procedures . . . . . . . . . . .
Sonication of Algae. . . . . . . . . . . . .
Enzyme Assays. . . . . . . . . . . . . .
Extraction and Determination of Chlorophyll.
RESULTS AND DISCUSSION. . . . . . . .

Rate of Glycolate Formation in Tobacco
Enzymes in Algae . . . . . . . . . . .
Glycolate Oxidase. . . . . .
Isocitrate Dehydrogenase . .
NADH:Glyoxylate Heductase. .
PhOSphoglycolaifi Phosphatase
Distribution of C in Products of he
Glycolate Pathway Formed During l 002
Photosynthesis by Algae. . . lh°
The Effect ofcx-hydroxysulfonateso on 002
Photosynthesis by Algae. . . .
Environmental Factors Affecting .Glycolateo
Production and Utilization by Algae. . . .
Growth and Photosynthesis of Algae in
Bed and Blue Light . . . . .
Manganese Deficient Culture of Chlorella
Glycolate Uptake and Excretion . . . . .

O O O O O O O
O O 9 0 0 0 O

O
0
O
0
O
O
0

SUMMARY 0 O O O O O O 0 0 0 O O O O O 0 O O 0 O 0
BIBLIOGRAPHIC O O O 0 O 0 O O 0 O O O 0 O 0 0 O 0
APPENDIX 0 O O 0 O 0 O O O 0 O O O O O 0 0 0 0 0 0

0 O O O O 0 O 0

75
78
121

121
154
159
167
174

186

LIST OF TABLES
Table No.

l Photosynthesis experiment with tobacco
leaves 0 O 0 0 O O O O O O O O O O O O 0 O O 32

2 Percent distribution of 1“c in standard
03c°mp°undseeeeeeeeeeeeeeee "’1

3 Percent distribution of lac in standard 2
czcompoundseeeeoeeeeeeeeeee “'2

h pH of algal suspensions before and after
aonication. O O O O O O O I O O O O O O O O M

5 Specific activities of compounds isolated
fromutobacco leaves after photosynthesis. .
COZeeeeeeeeoeeeeeeeee 61

6 Percent distribution of 1“c in 3-phospho-
glycerate isolated from tgbacco leaves
after photosynthesis in1 002 . . . . . . . 64

7 Percent distribution of 140 in glycerate
isolated from tobaifio leaves after '
photosynthesis in C02 .. ... . . . . . . 65

8 Percent distribution of 1“C in glycolate
isolated from tobacco leaves after
photosynthesis in 002 . . . . . . . . . . 66

9 Specific activities of carbon atoms in PGA.
glycerate and glycolate which were
isolated from tobaigo leaves after
photosynth6818 in 002 e e e e e e e e e e 68

10 A summary: Preparation of cell-free
extracts from algae and the detection of
enzymes in these extracts . . . . . . . . . 7“

11 Percent distribution of 1“c in the carbons
of phosphoglycerate, glycine and serine
formed'by algae during photosynthesis . . . 76

12 Chlorophyll content of cultures adapted to
blue light 0 O O O O O O O O O O O O O O O 0 127

13 Chlorophyll content of cultures adapted to
blue light 0 I O O O O O O O O O O O O O O 0 128

iv

Table No.

lu Percent distribution of 140 fixed by
Chlorella adapted to blue light a ter
3 and 10 minutes in the dark in 1 002. . . . 1“?

15 Percent distribution of 140 fixed by a
0.5 % suspension of Chlorella (grown
in white light) during photosynthesis
in blue light (“00-500 mu) . . . . . . . . . 150

16 Percent distribution of 1”0 fixed by a
0.5 % suSpension of Chlorella (adapted
to blue light) during photosynthesis
in blue light (#00-500 mp) . . . . . . . . . 151

17 Percent distribution of 140 fixed by a
0.5 % suSpension of Chlamydomonas
(adapted to blue light) during photo-
synthesis in blue light (400-500 mp) . . . . 152

18 Percent distribution of 140 fixed by
Chlorella pyrenoidosa (Chick). . . . . . . . 158

19 Glycolate excretion by Chlorella ,
pyrenoidosa (Chick). . . . . . . . . . . . . 160

 

20 Uptake of glycilate-Z-luc or phospho-
glycolate-Z- C by Chlamydomonas
reinhardtii in white light . . . . . . . . . 161

 

21 Excretion and uptake of glycolate by

Chlorella pyrenoidosa (Chick). . . . . . , , 162
22 Percent distribfition of 180 after feeding
glycolate-l-l C. 15“

10

ll

12

13

APPENDIX TABLES

Percent distribution of 1LLC fixed by

tobacco leaves. . . .

0 0 O O O O 0 0

Percent distribution of lac fixed by

tobacco leaves. . . .

Percent distribution of

Chlamydomonas at pH 7.

1“0 fixed by

6 O 0 0 O O 0 0

Percent distribution of luC fixed by

Chlamydomonas at pH 6.

5 . . . . . . .

Percent distribution of 1“c fixed by

Chlamydomonas at pH 8.

Percent distribution of

Chlamydomonas at pH 6.

Percent distribution of

Chlamydomonas at pH 8.

3 . . . . . . .

luc fixed by

5 . . . . . . .

luc fixed by

l O O O O O 0 0

Percent distribution of luc fixed by
Chlamydomonas during initial days of
growth in blue light (400-500 mu) . .

Percent distribution of

140 fixed by

Chlorella during initial days of
growth in blue light (400-500 mu) . .

Percent distribution of
Chlamydomonas adapted
(900-500 mp). . . . .

Percent distribution of
Chlamydomonas adapted
(400-500 mu). . . . .

Percent distribution of
Chlamydomonas adapted
(>600 mu) . . . . . .

Percent distribution of

1(“’0 fixed by
to blue light

I O O O O O 0 O

140 fixed by
to blue light

140 fixed by
to red light

0 O O O 0 0 0 0

Inc fixed by a

0.5 % suspension of Chlorella (grown
in white light) during photosynthesis
in blue light (“00-500 mp.)o o o o e o

vi

186

187

188

189

190

191

192

193

194

195

196

197

198

19

15

l6

17

18

Percent distribution of 140 fixed by a
0.5 % SuSpension of Chlorella (grown
in white light) during photosynthesis
in blue light (900-500 mu). . . . . . .

Percent distribution of 140 fixed by a
0.5 Z suSpension of Chlorella (adapted
to blue light) during photosynthesis
in blue light (400-500 mu). . . . . . .

Percent distribution of 180 fixed by a
0.5 % suSpension of Chlorella (adapted
to blue light) during photosynthesis
in blue light (900-500 mp). . . . . . .

Percent distribution of 1”c fixed by a
0.5 S suspension of Chlamydomonas
(adapted to blue light) during photo-
synthesis in blue light (400-500 mu). .

Percent distribution of 1&0 fixed by a
0.5 % suSpension of Chlamydomonas
(adapted to blue light) during photo-
synthesis in blue light (400-500 mp). .

vii

199

200

201

202

203

LIST OF FIGURES

Figure Title Page
1 The glycolate pathway and its relationship
with COZ fixation. O 0 O O O O .1 O O O O O 3
2 Absorption spectra of filter systems used
for algal culture and_photosynthesis
enerimen‘tscee...oet....... 27
3 Total fixation of lucoz by tobacco leaves. . 51

Distribution of 1“c in PGA. glycerate.and
glycolate formed by tobacco leaves . . . . 53

5 Distribution of lac in glycolate, glycine,
serine. and sucrose formed by tobacco‘ 6
leaves 0 O O O O O O O O O O 0 O O O O O O 5
6 Total 1“c in PGA. glycerate. and glycolate 8
formed by tobacco leaves . . . . . . . . . 5
7 Specific activities of PGA. glycerate. and

glycolate formed by tobacco leaves . . . . 62

8 Badioautographs of methanol-water extract
from tobacco leaves after 20 and 300
secondsphotosynthesis in 1 002 in full
sunlighteeeeoeeoeoeeeeeeo 80

9 Radioautographs of methanol-water extract
from.Chlamydomonas-:fter 60 seconds
photosynthesis in 1 002 and 3000 ft-c,
'hit.118hteeeeeeoeooeooeec 82

10 Total fixation of 1“co by Chlamydomonas
reinhardtii at pH 7.6. . . . . . . . . . . 84

11 Percent distribution of 1""0 in some pro-
ducts in the methanol-water soluble
fraction formed by Chlamyggmonas
during photosynthesis in 002 . . . . . . 86

12 Percent distribution of 1&0 in some pro-
ducts in the methanolewater soluble
formed by tobacco i Ves during
photosynthesis in 002. . . . . . . . . . 88

viii

13

1L1

15

l6

17

18

19

20

21

22

23

24

25

26

Total fixation of 1“'CO by Chlamydomonas
reinhardtii at pH 6.5 . .

O 0 O O O O 0 0

 

Total fixation of 1400 by Chlamydomonas
reinhardtii at pH 8.; . .

 

Percent distribution of 140 in PGA.
glycolate.and amino acids formed by
Chlamydomonas at pH 6.5 . . . . . . . . .

Percent distribution of 140 in PGA.
glycolate, and amino acids formed by
Chlamydomonas at pH 8.3 . . . . . . . . .

Total 140 in ribulose-l.5-diphosphate
formed by Chlamydomonas . . . . . . . . .

Total fixation of 14co by Chlamydomonas
reinhardtii at pH 6.8

O O O O O O O O O 0

 

Total fixation of 11+C02 by Chlamydomonas
reinhardtii at pH 8.1 . . . . . . . . . .

 

Percent distribution of 140 in PGA,
glycolate, and amino acids formed by
Chlamydomonas at pH 6.5 . . . . . . . . .

Percent distribution of 140 in PGA,
glycolate. and amino acids formed by
Chlamydomonas at pH 8.1 . . . . . . . . .

Total 1“'0 in ribulose-l.5-diphOSphate
formed by Chlamydomonas . . . . . . . . .

Percent and total 11+C in ribulose-l,5-
diphosphate formed by Chlamydomonas . . .

Growth rates of a Chlamydomonas reinhardtii
culture in blue light (4004500 mu). . . .

Growth rates of a Chlorella pyrenoidosa
culture in blue light (400-500 mu). . . .

ChlorOphyll a/b ratio in green algae during
growth in blue light (400-500 mp) . . . .

ix

92

94

96

98

101

103

105

108

110

112

114

122

124

129

27

28

29

30

31

32

33

34

In vivo Spectra of Chlamydomonas

—reinhardtii grown in blue light

 

(400- 500 mu). . . . . .

Percent distribution of 140 in products
formed by Chlamydomonas reinhardtii
after adaptation to colored light during

photosynthesis in l1‘1C02 . .

Total 1“C in glycolate, glycine, and
serine formed by Chlamydomonas reinhardtii
adapted to red light (>600 mu) during

photosynthesis in1C02 . .

Total 140 in glycolate, glycine.and serine
formed by Chlamydomonas reinhardtii

O

 

adapted to blue liggt (400- 500 mu7—during

photosynthesis in 002 . .

Total Inc in TCA cycle acids formed by
Chlamydomonas reinhardtii adapted to red
I4 blue light during photosynthesis in

C

 

O

0

O

0

O

0

OZeoeoeeo.eeooeeoeooe

Total 11+C in sugar phosphates by

Chlamydomonas reinhardtii adapted to
either red or Blue light during photo-

synthesis in

Total lac fixed by green algae during
photosynthesis in various in ensities of

blue light (400-500 mu) in 1

Growth rates for manganese deficient

C02.

0

C02 0 e e o o o o o o o o o

cultures of Chlorella pyrenoidosa (Chick)

 

during growth in white light.

132

135

138

140

142

144

148

156

Be
Pun
ft-c
a-xc
m
NADR
NADP
NADPH

dePO
ORSHSulfonate
OHAPHSulfonate

PGA
P-Glycolate
POPOP

PPO

PS

RuDP

S.A.

Sugar-P

TCA cycle
UDPG

LIST OF ABBREVIATIONS

Pyridoxal PhoSphate

Flavinmononucleotide

foot candles

«bketoglutarate

Nicotinamide-Adenine Dinucleotide
Nicotinamide—Adenine Dinucleotide (Reduced)
Nicotinamide-Adenine Dinucleotide Phosphate

Nicotinamide-Adenine Dinucleotide Phosphate
(Reduced) ‘

d-naphthylphenyloxazole
azhydroxymethanesulfonate

.uhhydroxypyridinemethanesulfonate or
2-pyridylhydroxymethanesulfonate

3-Ph08phoglycerate

. Phosphoglycolate

l.4-bis-2-(5-phenyloxazolyl)benzene
2.5-diphenyloxazole

Photosynthesis
Ribulose-l.5-diphosphate

Specific Activity

Sugar PhoSphates

Trichloroacetic Acid

Tricarboxylic Acid Cycle

Uridine Diphosphate Glucose

xi

INTRODUCTION

The "path of carbon in photosynthesis" describes how
photosynthetic assimilatory power. ATP and NADPH produced
in photosynthetic electron transport. is utilized for
carbon dioxide fixation into the organic compounds of green
plants, algae. and photosynthetic bacteria. Kinetic iso-
tope tracer studies, initially develOped for photosynthesis
by Calvin's group, enabled a determination of the sequential
formation of compounds during an isotOpe feeding. During
early investigations carboxyl labeled 3-phosphoglycerate was
identified as the initial product of photosynthetic 11+002
fixation. Subsequent metabolism of 3-phosphoglycerate
successfully described the regeneration of the 002 acceptor,
ribulose-l,5-diphosphate, through a sequence of reactions
involving sugar mono- and diphosphate esters of both the
Embden-Meyerhoff glycolytic pathway and the pentose phosphate
pathway (9).

Many publications on the enzymes involved in this path-
way and on the physiological significance of the carbon cycle
have confirmed the initial pr0posals of Calvin's group. This
study discusses some observations of 002 fixation which can-
not be explained by that cycle. namely, the very rapid
incorporation of 1”002 during photosynthesis into the compo-

nents of the "glycolate pathway": phosphoglycolate,

2

glycolate, glyoxylate, glycine, serine, glycerate. and
sucrose. The labeling of these products, which may vary
from 10 f to 90 % of the total 1""002 fixed, depends upon
many physiological conditions and the length of the ex-
periment. These compounds of the glycolate pathway together
with the compounds of the photosynthetic carbon cycle account
for nearly all of the soluble 140 products of short term
1"1002 photosynthesis.

The importance of the two carbon acid, glycolic orcx-
hydroxyacetic acid, in photosynthesis remains obscure.

Although rapidly labeled during 1“

C02 fixation, glycolate's
synthesis has never been directly implicated in photo-
synthetic 002 fixation. Several laboratories, investigating
the metabolic role of this acid in higher plants (47,90,120)
and algae (88,89,125), have develOped the glycolate pathway
to describe the metabolism of this acid. A schematic
representation of this pathway and its relationship to the
carbon cycle is shown in Figure l.

The photosynthetic research of 0. Warburg has also
suggested an important role for glycolate. Warburg has re-
ported that up to 90 i of the C02 fixed during photosynthesis
by his Chlorella is present as glycolate under certain
conditions (121). From low quantum yields for photosynthesis

in Chlorella warburg has suggested that the additional energy

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required for the reduction of C02 to carbohydrate is pro-
duced by a "Rfickreaction" (122). We wonder whether this
energy requirement might be obtained from the reoxidation of
reduced carbohydrate components to glycolate or even the
further oxidation of glycolate to glyoxylate.

This investigation evaluates the biosynthetic condi—
tions and catabolic fates of glycolate in higher plants and
algae. In both higher plants and algae, good comparative
data has been lacking for the rate of 1“c labeling of each
carbon atom of phOSphoglycerate, glycolate, glycine, and
serine. Data in previous publications (87,88,89,104,107,
125) have been interpreted assuming that the glycolate path-
way does function in green algae, although no direct
evidence supporting this assumption was presented. As re-
ported in this thesis, the absence of glycolate oxidase and
the glycolate pathway in algae requires a reevaluation of

these previous experiments.

LITERATURE REVIEW

Glycolate Biogynthesis:

Benson and Calvin (l3) and Wilson and Calvin (127) have
reported the rapid labeling of glycolate by algae during
photosynthesis; uniformly labeled glycolate was reported by
Schou,et_alg (95). The kinetics of 1&0 accumulation in
glycolate during photosynthesis (127) demonstrated that gly-
colate was likely a product of the carbon reduction cycle
intermediates. A possible source of uniformly labeled
glycolate is the sugar phosphates which when cleaved and
oxidized would produce glycolate. Griffith and Byerrum (32)
reported a conversion of ribose-l-luc to glycolate in tobacco
leaves although yields of glycolate-lac were quite low. The
formation of glycine from ribose-l-luc has been demonstrated
by Weissbach and Horecker (123) in which they implicated the
intermediates glycolate and glyoxylate. Zelitch.(l38) has
reported a 10 Z conversion of uniformly labeled ribulose-luce
phosphate to glycolate by Spinach chlor0p1asts in the presence
of an.«-hydroxysulfonate.

Using the above arguments suggesting that glycolate
could be formed from the sugar phoSphates of the carbon re-
duction cycle, Bassham has created a hypothetical scheme for
glycolate formation involving thiamine pyr0phosphate (TPP)
and other classical 02-metabolites (8). He presented no new

experimental evidence, however, to support this concept.

6

7

although it has been suggested by data from Holzer's group.
Using TPP and pyruvate oxidase or transketolase they
isolated a glycolaldehyde-TPP intermediate (28). Later, in
agreement with reports from Hacker's laboratory (15,31),
Holzer found that a glycolaldehyde-TPP intermediate formed
from a pentose or hexose phosphate could be oxidized to
glycolyl-TPP in the presence of ferricyanide which then was
hydrolyzed to release free glycolate (43). The in 3112
significance of these reactions is unknown, but these ex-
periments establish a background for the hypothesis that
glycolate might be formed via a cleavage of a sugar phosphate
to a TPP-CZ complex which in turn is oxidized to free
glycolate.

Richardson and Tolbert (92) have isolated and partially
purified a Specific phOSphatase for phosphoglycolate. Iu,§t_
_al; (131) have also found this enzyme in Spinach and wheat
leaves. Recently Bassham and Kirk (11) have confirmed the
rapid labeling of glycolate by Chlorella and also reported
that 02 stimulated the production of both glycolate and phos-
phoglycolate. Tolbert (107) has suggested that phospho-
glycolate might be formed from the cleavage of one of the
sugar diphosphates presumably through a TPP-C2 intermediate.
The oxidation of this 02-phosphate fragment would produce the
phosphoglycolate which phOSphoglycolate phosphatase would

hydrolyze to free glycolate. The inconsistent observation of

8

phosphoglycolate in photosynthesis experiments is probably a
reflection of the resistance of the Specific phOSphatase to
inactivation in methanol. Ullrich (116) has reported that
terminating experiments in methanol does not prevent some
hydrolysis of phOSphate esters, and particularly the hydro-
lysis of phosphoglycolate.

In spite of the evidence against glycolate as an initial
product of algal photosynthesis (81,127), Zelitch has re-
cently published that glycolate cannot come from a phospho-
glycerate metabolite (139). Stiller (101), in a recent
review has criticized classical arguments for PGA as the first
product of photosynthesis and discussed the possible gg’ggzg
synthesis of glycolate. Using data from long time photo-
synthesis experiments. Zelitch (139) reported that the
Specific activity of glycolate is higher than the carboxyl
carbon of phosphoglycerate; thus. he suggested that glycolate
originates from a compound initially formed by the condensa-
tion of two 002 molecules. This route to glycolate is
schematically represented in Figure l as a 002 + 002
condensation. In addition to the length of Zelitch's experi-
ments (2 and 5 minutes), he used a killing time of about 30
seconds in 12002 air. All of his reported experiments were
inhibited with a "glycolic acid oxidase" inhibitor.¢x-hydroxy-
pyridinemethanesulfonate (137). No short time kinetic

experiments with tobacco leaves have been reported for the

9
initial formation of glycolate. In such experiments one must
consider both the rate of 140 incorporation into products as
well as specific activity. A consideration of each fact,
independent from the other, could lead to a misinterpretation
of the results. The actual specific activity might be
diluted if different resevoirs contributed to a common pool,
e.g..extremely high specific activity of a compound within
the chlor0p1ast could be greatly reduced by a large cyt0plasmic
pool of the same compound. 0n the other hand, the rate at
which the Specific activity of a compound approaches the
specific activity of the 114C02 may reveal the relative sizes
of two different pools.

Work with mutants of Chlamydomonas reinhardtii, which
lack the normal carboxydismutase of ribulosecl,5-diphosphate,
did not have a light fixation pattern significantly different
from that of dark fixation (62). The 140 was distributed
mainly in the acids of dark fixation: malate, aSpartate, and
glutamate. Since no glycolate formation was reported, no
support was given to Zelitch's hypothesis. In the mutant
algae, glycolate and the products of its metabolism Should have
been present, if a distinct, autonomous 002 fixation pathway
existed which was different from the known carboxylation of
ribulose diphOSphate leading to the formation of phospho-

glycerate.

10

The reduction of glyoxylate could also serve as a source
of glycolate as shown in Figure 1. In 1955 the prOperties
and action of a NADH specific glyoxylate reductase were de-
scribed; this enzyme catalyzed the reduction of glyoxylate
(135). In 1962 a second glyoxylate reductase was found
associated with chlor0p1asts which preferentially reacted with
NADPH (144). Since this reaction is reversible only with
extremely high excesses of glycolate. the enzymatic reaction
could function as a synthetic route from glyoxylate to
glycolate. However. no direct evidence for the early forma-
tion of glyoxylate during photosynthesis has been reported.

Two major sources of glyoxylate might be the formation
from isocitrate in the glyoxylate cycle. described by

Kornberg (54) in Pseudomonas §R., or from the deamination of

 

glycine. The glyoxylate cycle exists in many organisms.
bacteria (50,54,83), algae (34), and higher plants (19.129).
The glyoxylate cycle is most active in tissues which are re-
quired to metabolize much fat and therefore, not very active
in the mature leaves of plants. Seeds usually have a very
active enzyme complement for this cycle (16.76.98). In the
algae and tetrahymena, these enzymes are induced only after
growth in acetate (45,63,102). Apparently, the normal magni-
tude of this cycle must be small in actively photosynthesizing
tissue. waever. further information is needed in order to
evaluate the low levels of isocitrate observed by Asada,et_all

(4) in tobacco leaves inhibited with<x-hydroxysulfonate. It

11

is unlikely that the turnover of this cycle can account for
either the early labeling of glycolate in photosynthesis. or
the distribution of 14C in the organic acids when glycolate
metabolism is inhibited. Therefore, this cycle should not be
considered as important as other postulated sources of
glycolate in higher plants.

The work of Homann (44) may also suggest another mechanism
for glycolate biosynthesis. Using a model system with 2,3-
diketogulonate, he proposed a possible oxidation mechanism
with Mn"'3 and the hydrated otfi-dicarbonyl moiety of the di-
ketogulonate which resulted in the cleavage and oxidation of
the 1,2 carbon fragment to oxalate with the concomitant re-
duction of Mn+3 to MnIZ. Such a mechanism might also describe
the formation of glycolate, since the requirement for Mn in
glycolate formation has been reported (103).

Glycolate could also be formed from glycolaldehyde.
Davies has reported that a NAD-dehydrogenase, specific for
glycolaldehyde was present in the mitochondria from etiolated
pea seedlings (24). However, a glycolaldehyde dehydrogenase
has not been reported elsewhere, although Tolbert's labora-

tory has looked for it (personal communication).

Glycine Biosynthesis:
Vernon and Aronoff reported that glycine is not formed
directly in photosynthesis (117). Tracer experiments with

glyoxylate demonstrated its direct conversion to glycine (49,

12

60.97.99.120). The production of glycine from glyoxylate has
been reported as a B6 transamination reaction in carrot, pea
leaves, and corn coleOptiles (22). In castor beans a trans-
aminase Specific for alanine has been reported by Kretovich
(57), and Morgunova has also observed a glyoxylate transaminase
in oil-bearing plants which preferentially utilizes f-amino-
butyric acid (73). The reductive amination of glyoxylate by
a soy bean fraction is a reaction which may be important in
nitrogen fixation (56), although this reaction has not been
confirmed elsewhere.

The production of glycine from serine and the formation
of "active-formaldehyde" using tetrahydrofolic acid by re-
versal of the serine hydroxymethylase could be another source

of glycine (67,69,85). When serine-la

C was fed to leaves,
however, little of the label appears in glycine but rather

the lac moves into glycerate and sugars (90,119.14). Although
the formation of glycine from serine has been reported in
wheat (69,118) and Chlorella (14), this conversion is limited
and does not likely function in Cl metabolism as readily as

in E. 921; (85). The inhibition studies of Whittingham and
Pritchard (125) presented evidence that the conversion of
glycine to serine does occur to a significant extent in algae,
but the general nature of isonicotinyl hydrazide inhibition

suggests that glycine formation in algae proceeds via some

pathway not described above.

l3

Serine Biosynthesis:

The formation of serine in organisms has been well docu-
mented. Serine synthesis from glycine is an important
reaction in plants as indicated by studies with formate and
glycolate metabolism. The formyltetrahydrofolic synthetase
has been isolated from higher plants as a cytoplasmic enzyme
and can account for the activation of the 01 unit (42).
Isotope feeding studies with higher plants demonstrated that
formate was incorporated into thegf-carbon of serine and
required tetrahydrofolic acid and B6 (23,35,105). The tetra-
hydrofolic acid requirement has also been observed in
mammalian tissue (52) and bacteria (93).

Evidence for serine formation from the transamination
of hydroxypyruvate has been reported in pea seedlings(58),
and an alanine:hydroxypyruvate transaminase has been de-
scribed in higher plants (126). The formation of serine from
PGA by algae or chlor0p1asts is supported by the labeling
distribution in PGA and serine (20,90,133). Phosphatase can
convert PGA to free glycerate (114) which can then be con-
verted to hydroxypyruvate by D-glycerate dehydrogenase (100).
and then the hydroxypyruvate transaminated to serine.
Alternatively, the production of serine from PGA has also been
postulated directly through phOSphohydroxypyruvate to phos-
phoserine as Shown in pea coleoptiles (33). Such a pathway

has also been observed in human cells (84).

14

In summary, serine biosynthesis most likely occurs by
one of the three pathways described above: the incorporation
of a C1 group activated by tetrahydrofolic acid into glycine
in the serine hydroxymethylase reaction, the transamination
of hydroxypyruvate formed from PGA and glycerate, or the
transamination of phosphohydroxypyruvate from PGA and the

hydrolysis of phOSphoserine to serine.

The Glycolate Pathway in Higher Plants:

The initial isolation and characterization of glycolate
oxidase by Tolbert,gt_a1; (108) formed the basis for subse-
quent work on glycolate metabolism. Glycolate oxidase
catalyzes the oxidation of glycolate to glyoxylate and H202
(Figure 1). This enzyme was further purified and characterized
(142) and finally crystallized in the presence of FMN by
Frigerio and Harbury (29). The oxidase has a molecular weight
of 70,000 and a pH Optimum of 8.3.

Glycolate oxidase is present in all higher green plant
material capable of photosynthesis (59). Recently, however,
Mitsui,gt‘al& (71,72) have reported that glycolate oxidase is
present in the root tips of low-land rice plants, and Tanner
and Beevers (103) have reported glycolate oxidase activity in
castor bean endosperm. Current investigations indicate that
an alternate form of the enzyme may exist which may be
associated with the electron carrier ferredoxin (6). Thus,

the oxidase may interact in a coupled electron tranSport

15

system, although this has not yet been demonstrated. The
oxidase activity increases as the plant becomes green. or the
oxidase can also be activated in etiolated tissue by the
addition of glycolate or flavin mononucleotide (59.109).

The importance of this oxidase in photosynthesis has been
inferred from the dependence of the enzyme's synthesis on
light. the early appearance of glycolate in photosynthesis,
and the lack of serine formation in etiolated wheat tissue
until the appearance of glycolate oxidase activity (111). An
investigation of the possible role of the oxidase in oxida-
tive_phosphorylation was negative (141). so that further
investigations concentrated on the role that the glycolate
oxidase played in the metabolism of organic acids in higher
plants.

The utilization of glycolate by plant materials has been
extensively investigated and all bf the data seem to be con-
sistent with the pathway as shown in Figure l. Talbert and
Cohen (110) in feeding higher plants labeled glycolate-Z-lnc
found that glycine and serine were the main metabolic products
of glycolate and that a( and ,6 carbons of serine were uniformly
labeled. These initial studies were later complemented with
more data on the distribution of the labeled carbon in the
compounds formed from glycolate. Rabson, Talbert and Kearney
(90) confirmed the above results and extended the sequence of

reactions to glycerate. using additional data from serine-3-lu0

l6

feeding and 11+COthotosynthesis. The data of Wang and
Waygood (120) also indicate that a glyoxylate-serine pathway
is present in higher plants. The 11+0 distribution pattern

in the sugars after the feeding of labeled serine proved that
a sugar reservoir in wheat or soybean leaves could be formed
from these precursors (47,90,120). A glycerate kinase as
reported by Ozaki and Wetter (82) was found in rapeseed to
function in the conversion of glycerate to phosphoglycerate.
The hexoses of sucrose were labeled in the 3,4 positions

1”C, and in the 1,6 positions after

after feeding serine-l-
feeding serine-3-luC to the plants. The main pool of

labeled glycerate, however, formed by the soybean (3) or

wheat (90) is not uniformly labeled as eXpected if synthesized
by the glycolate pathway.

To evaluate whether or not the glycolate pathway is
associated with the cyt0plasm or the chloroplast, the dis-
tribution of 140 in serine formed by isolated chlor0p1asts
was compared with that from the whole spinach leaves after
photosynthesis in lI‘I'COZ (20). Carboxyl labeled serine was
found in the chlor0p1ast while that from the whole plant was
uniformly labeled. These results were interpreted to indicate
that two pathways exist for serine biosynthesis, but that
the glycolate pathway was by far the most active since the

serine pool from the whole plant was uniformly labeled after

20 seconds photosynthesis (90.95).

17

Other evidence which supports the metabolic function of
the glycolate pathway comes from the use of inhibitors.
Zelitch (134,137,143) and Asada,§t_a14 (4) have usech-hydroxy-
sulfonate inhibitors which Zelitch had described as specific
for glycolic acid oxidase (134). These metabolic inhibitors
have been used to evaluate the in 1132 Significance of the
glycolate pathway, and have revealed that as much as 50 % or-
more of the fixed carbon accumulates in glycolate. Thus, much
of the carbon dioxide fixed during photosynthesis must be
channeled through this pathway.

The Glycolate Pathgay in Algae:

Schou, _e_1; if. (95) fed glycolate-1-1% and glycolate-Z-luC
to Scenedesmus and reported that glycolate was metabolized
into amino acids and sugar phOSphates in the light with equal
labeling in the O‘and.6 positions of glycerate. This dis-
tribution of 1"(C in glycerate is consistent with its being
formed via the glycolate pathway typical of higher plants.
However, Smith,et,a1, (99) suggested that in algae both
alanine and serine were formed from PGA. The presence of the
glycolate pathway enzymes in algae has not been well docu-
mented. In a preliminary report of part of this work (40),

we have shown the absence of glycolate oxidase in three genera
of algae. A report by Kolesnikov (53) and another by Ohmann
(78) have also suggested that glycolate oxidase, if present in

algae, is probably different from the higher plant oxidase.

18

Also. in an initial report Tolberu gt a1; (112) noted that in
Chlorella and Chlamydomonas thecx-hydroxysulfonate inhibitors
described by Zelitch (134,137) did not cause an accumulation

of glycolate during 1"

002 photosynthesis as in the higher
green plants.

Whittingham's laboratory has used another inhibitor, iso-
nicotinyihydrazide which caused the accumulation of glycolate
and glycine during photosynthesis with Chlorella (87.88.89.
124,125). This inhibitor has been shown to inhibit pyridoxyl
phosphate (36) enzymes (130) and therefore inhibits the
possible conversion of glycine to serine in the glycolate
pathway. Whittingham explains the accumulation of 140 in
glycolate and glycine. but never considers that inhibitors of
B6 enzymes should also decrease glycine accumulation since
glycine too is likely formed via a pyridoxal phOSphate
mechanism (55).

In a Russian paper, Zak and Nichiporovich (133) reported
the 170 distribution in glycine, serine. and alanine after
photosynthesis by Chlorella in 14002. He found that alanine
and serine were labeled similarly to PGA; and the labeling
pattern of glycine excluded the possibility of its formation
from serine. Vernon and Aronoff (117) reported that alanine
and serine formed in soybean leaves.but only alanine was
labeled similarly to PGA. This data and that outlined above
suggest that the glycolate pathway may not exist in algae as
it does in higher plants.

19

Glycolate Excgetion and Uptake:

Tolbert and 2111 (113) reported that free glycolate was
excreted by Chlorella into the algal medium during photo-
synthesis. Later it was also shown that chloroplasts
specifically excrete glycolate (49,106). since the photo-
synthetate found in the supernate was almost exclusively
glycolate. These observations of excretion have been con-
firmed by Pritchard,gt.a1L (87). Millen gt_alg (70). Fogg.gt
a1; (26.27), and Nalewajko,§t_alg (74). and recently such
excretion has been reported for Chlamydomonas and Ankistro-
desmus (40). The conditions for glycolate excretion are
similar to those for glycolate production: high conditions
of pH. light intensity and oxygen concentration; the presence
-of bicarbonate ion or low 002 concentration; and the presence
of manganese.

As stated above. Schou.et a1. (95) observed some
glycolate-170 could be converted by Scenedesmus at pH 2.8
in the light into amino acids and sugar phosphates. although
no data was given for the amount of glycolate utilized.

Fogg has develOped a theory which considers the excreted
glycolate as a reservoir of chemical energy, accumulated
during photosynthesis. which can be later used by the phyto-
plankton. On the other hand Kearney and Tolbert (49) have

reported that glyoxylate and not glycolate is taken up by

20

chloroplasts. The small quantities of glycolate uptake.
under a variety of conditions, suggest limited metabolism of
this acid (40). which is in contrast to acetate uptake and

utilization by green algae (77).

Function of the Glycolate Pathway:

There have been several postulated functions for the
glycolate pathway in higher plants (10?). The excretion of
glycolate by chlor0p1asts (106) and the cytOplasmic function
of glycolate oxidase may Operate together in tranSporting
the fixed carbon from the chloroplast into the metabolic
pools of the cyt0plasm. Ongun and Stocking (79,80) have
recently reported that the chloroplast membrane is freely
permeable to serine and thus serine can function in a trans-
port mechanism between the chlor0plast and the cyt0plasm.
Their hypothesis, however. lacks any directional prOperty
which is required in a system expressing net transport.
Their conclusions are also not supported by the recent evi-
dence of Chang and Tolbert (20) that serine in the chlorom
plasts is labeled differently from that of the whole plant.

That algae also excrete glycolate (113) suggests that
glycolate might be involved in an anion pumping mechanism
e.g.,for bicarbonate transport (106). The stimulation of
glycolate production in low 002 concentration is consistent
with the anion pumping mechanism. Since at higher concentrations

of 002 transport need not be dependent upon glycolate production.

21

Tolbert (107) has also suggested that the pathway might
function in a permease system which is given direction by
phosphoglycolate phOSphatase. The phOSphoglycolate may be
hydrolyzed on the membrane and released as free glycolate
which cannot reenter the chloroplast (49). Although many
hypothetical reactions for the production of energy from
glycolaldehyde etc. have been suggested,there has been no
evidence for the coupling of these enzymes with electron
transport. The production of glycolate might also be involved
in the 02 evolution steps Of photosynthesis. since both
processes are sensitive to CMU (107) and require Mn (103).

Since glycolate oxidase is cyanide insensitive (108) and
very active in higher plants (21). the oxidation of glycolate
may be involved in the reSpiration of leaves as a terminal
oxidase (21). Based on both the amount of enzyme and the
amount of glycolate in higher plants. the oxidation of
glycolate could account for the total 02 uptake of tobacco
leaves (136,137,140).

The glycolate pathway might also be involved in the
control of stomatal Opening (143). Zelitch proposed that
synthesis of sugars from glycolate via the glycolate pathway
alters the osmotic potential of the guard cells. Glyoxylate
reduction via glyoxylate reductase might function as a sink
for excess reducing power produced as NADPH in photosynthesis.

which if formed in large excess could inhibit photosynthetic

22

electron tranSport and phosphorylation. The oxidation of
NADPH would favor the formation of ATP which may be required
for stomatal opening. Both Heath,et_a14 (39) and Mansfield
(65) have suggested that both of these hypotheses are in—
correct.

A preliminary report suggested that glycolate oxidase
might also function in glucose uptake (17). In an argument
similar to that of Zelitch described above, the authors
suggested that glyoxylate reduction could eliminate excess
NADPH and therefore stimulate ATP production in photo-
synthesis. Since their only data was a correlation of glucose
uptake andud-hydroxysulfonate inhibition of this uptake,the
implication of glycolate metabolism in glucose tranSport is

probably a misleading interpretation of results.

Environmental Factors Affecting Glycolate Formation:

The action spectrum for photosynthesis has long been .
recognized to be approximately that Of chlorOphyll a
absorption. although the careful work of many investigators
on responses of the initial photosynthetic events to different
light quality has implicated two interacting pigment systems
in photosynthesis. In green plants the two major pigment
systems contain chlorophyll a and chlorOphyll b. The
dependence of the relative concentrations of related pigments
on light quality has been reported. It has been difficult to

evaluate whether the changes in pigment concentrations were

23

caused by the reduction of light intensity (91) or whether
the quality of the light also induced these changes. The
recent work of Fujita and Rattori with Tolypothrix indicated
that changes in pigment ratios reSponded to a change in
light quality rather than light intensity (30). Jones and
Meyers have also completed similar work with Anacystis and
conclude: "There is a preferential increase in the pigment
of highest absorption for wavelengths of highest intensity"
(48). These results clarify the role of light quality in
pigment formation. while the light intensity apparently
affects the rate of pigment conversions (48).

In 1955, Roux.gt a1, (94) and Tyszkiewicz (115) reported
that plants. exposed to light with the emission spectrum of
chlorOphyll absorption. synthesized PGA and other sugar
phosphates. waever. if the incident light quality was
limited to the region of the spectrum absorbed by carotenes,
mainly amino acids were formed and no starch synthesis was
observed. Three reports (Rauschild.et_a1. 36.37.38) impli-
cated the interaction of red and blue light in 002 fixation
by both green and bluefgreen algae. In their experiments
blue light enhanced amino acid synthesis and decreased 140
accumulation in sugar phOSphates. These were long time (30
min) experiments. however. which had been preceded by a long
period in the dark (up to 3 hr). Another recent report from
Russia indicated that Chlorella in blue light fixed more 1“0

24

into the amino acids of the glycolate pathway and glycolate,
especially in the presence of increased 02 concentrations
(132). This result is in agreement with a preliminary report
of the work in this thesis (41). Likewise, Andreeva and
Korzheva (1) have reported that less alanine, but more
aSpartate, serine, and glycine were formed in sunflower
leaves if the plants were cultivated with blue light or if
the photosynthesis experiment was performed in blue light.

Recently Horvath and Sazasz have shown that the light
intensity can also affect the metabolism of the photo—
synthetic products (46). Their results confirm the vast
amount Of literature indicating that more amino acids are
produced in low intensity white light and that in high in-
tensities, starch is the main product. These results seem
consistent with previous work and theories of photosynthetic
production of ATP and NADPH.

That enzymes may respond to light has been shown in
several cases. The accumulation of glycolate oxidase in the
light has been discussed (59). The work of Margeulies on
the formation of glyceraldehyde dehydrogenase in red light
(66) may be relevant since this relates to photosynthetic

requirements and metabolism in the cell.

en—

METHODS
Plant Material:
Tobacco:

The tobacco plants used in these experiments, tobacco
(var. Maryland Mammouth), were sown May 18, 1965 in the green-
house and transplanted to an open field on June 11. Mature.
but not fully expanded, leaves were harvested for the
photosynthesis experiment July 21. Since the weather had been

quite dry, the plants were only 45-55 cm tall.

Algae:

The strains of algae used in these eXperiments were
obtained from the ”Culture Collection of Algae” at Indiana
University, Bloomington. Ind. Ankistrodesmus braunii (Naeg)
Collins (#245); QElQIEllE.EII§EQlQQ§a.(#395) Chick, Emerson;
and Scenedesmus gpliguus (#393) Krager. Gaffron D-3, were

cultured on an inorganic salt medium with Hoagland's microm
nutrients (75). Chlamydomonas reinhardtii (#90) Dangeard (-)
strain grew well on the high phosphate medium described by
Orth et al. (81). The cells were cultured in 2.5 liter "10w
form” Fernbach flasks fitted with air inlets; these were
placed on a reciprocating shaker of about 60 excursions per
min.. thus providing a gentle but thorough agitation of the
1500 ml of medium. For culturing of algae in white light
the shaker was held in a controlled environment chamber

(Sherer-Gillet) at 15°C which maintained a temperature in the

25

26

culture medium of 20°. The chambers provided 1200 ft-c of
continuous light from Westinghouse cool white super high
fluorescent bulbs (F96T12/CW/5HO). For aeration.and gassing
of the medium. air from an oil-free compressor was passed
through a cotton filter and distilled water and then mixed
with 002 (also bubbling through distilled water) so that a
final concentration of 0.25 C02 v/v in air was obtained.

For the culture of cells in portions of the Spectrum.
either blue and red filter systems were used as shown in
Figure 2. The red filter was PFire Red” #110 gelatin
(Grand Stage and Lighting 00.. Chicago. 111.). and light was
obtained from 15 watt red fluorescent tubes (General Electric -
FlSTlZ-R). This combination effectively eliminated all
radiation below 600 mp. Blue light was obtained from 15
watt blue fluorescent lights (General Electric FlSle-B or
Sylvania Fl5T8-B) which was filtered through a blue celluloid .
(supplied by Kliegel Bros. as medium blue #32 Cinemoid
filter) and 5 cm of a CuSOn solution of 30 g per 1. The
combination of blue celluloid and CuSOh solution produced a
rather narrow band of emission between 400 mp and 520 mp. All
of the culture work and photosynthesis experiments in blue or
red light were performed in dark rooms. Intensities of 350-
400 ft-o in the blue light and 200 ft—c in red light were used;
These cultures were grown under continuous light and aeration

with 0.2; 002 in air. at 20.210 as maintained by a water bath.

27

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29

Chemicals:

 

All of the chemicals used in this work were standard
reagent grade. The glyoxylate and hydroxypyruvate were
obtained from Nutritional Biochemicals (Cleveland, Ohio). and
phosphoglycolate as the cyclohexylammonium salt from General
Biochemical Co. (Chagrin Falls, Ohio). Chromatographically
pure sodium glycolate was produced by Matheson, Coleman and
Bell 00. FMN and the pyridine nucleotides, NAD, NADH, and
NADPH, were products of Sigma Chemical Co. (St. Louis, MO.):
NADP and isocitrate came from Calbiochem 00. (Los Angeles,
Calif.). Eastman Kodak (Rochester. N.Y.) supplied 2,7-
dihydroxynaphthalene (#4408) and chromotrOpic acid (#P230).

The inhibitor, 2-pyridy1hydroxymethanesulfonate (OH-PMSulfonate),
was obtained from Aldrich Chemical 00. (Milwaukee, Wis.L
N. E. Tolbert prepared the hydroxymethanesulfonate.

The counting systems utilized the following reagents: pa
dioxane (spectroquality) Matheson, Coleman and Bell: phenyla
ethylamine. BP 75-760/6mm. (#2642) Eastman Kodak (Rochester,
N.Y.): PPO (2.5-diphenyloxazole),cx-NPO Qx-naphthylphenyl-
oxazole), and POPOP (l,4-bis-2-(5-phenyloxazolyl)benzene) all
Scintillation grade, Packard Instruments (Downers Grove, 111.).

The Ba1#C03 received from Oak Ridge National Lab. was used
routinely in our photosynthesis experiments. The source of
uniformly labeled 3-phOSphoglycerate (1.35 mc/mmole) was
Calbiochem 00.. while uniformly labeled glycolate was prepared

30

and isolated from long time photosynthesis experiments in
11“002 with plant tissue. The following radioactive compounds
were purchased from Nuclear Research Chemicals. Inc.

(Orlando, Fla.): glycine-l-luc (3.4 mc/mmole). glycineuz-

11+C (2.0 mc/mmole). DL-serine-l-lnc (4 mc/mmole). DL—serine-3-
140 (6 mc/mmole). glycolate-Z-luc (4 mc/mmole), glycolate—la
170 (0.5 mc/mmole). and phOSphoglycolate-Z-luc (2.5 mc/mmole)°

Photppynthesis Experiment with Tobacco Leaves:

Young leaves from field grown tobacco (var. Maryland
Mammouth) were cut from plants in the morning and immediately
placed in a beaker containing either water or 0.01 M 2a
pyridylhydroxymethanesulfonate (OHAPMSulfonate) at pH 3.5.
These leaves were removed at 5 minute intervals so that each
leaf remained in solution exactly 1 hour before the photon
synthesis experiment. Partial shade and good ventilation in-
sured high transpiration rates so that the leaves took up
enough solution to become turgid in 10 minutes.

After 1 hour. the leaf was removed from the solution and
the blade tip cut so that the remaining leaf was about 15 cm
long and 10 cm across at the widest point. The tissue was
immediately put in the apparatus which had been designed for
rapid equilibration of the atmosphere with added 002 and for a
rapid killing procedure. The reaction chamber (leaknproof)

was a plexiglass box with internal dimensions of 11 cm x 15 cm

31

x 1.7 cm. The removable cover had two holes. 1.5 cm in diameter;
one held a stopper with a stopcock attached to a vacuum pump,
the other held a stopper with a 60 ml separatory funnel. This
separatory funnel contained a known amount of BalnCO3. and 2 ml
of lactic acid was introduced through a syringe cap on the tOp
of the funnel before the experiment was begun. A partial vacuum
of 75-90 mm Hg was pulled on the chamber and the vacuum line
closed. Then, at zero time, the stopcock of the funnel con»
taining the liberated 14002 was Opened and nearly simultaneously
the syringe cap removed by a second Operator so that the 11+002
was swept into the chamber. The leaf was held perpendicular

to the sunlight of an intensity of 7,000 ftmc as measured with
a Weston photometer. At the end of the experimental period the
lid of the chamber was slid Off while a second operator poured
hot absolute methanol into the chamber. Less than 2 seconds
were required to remove the lid and add the methanol. The
methanol and killed tissue were rapidly transferred to a beaker
and boiled for 5 minutes. After decanting the MeOH. the leaves
were boiled in water 2 minutes. The extracts were combined and
aliquots counted in the scintillation counter. Two-dimensional
chromatography (12) of these extracts permitted evaluation of
the product distribution of the fixed 1I‘l’C, while column chroma=
tography permitted isolation of larger quantities of several
compounds (139); The data in Table 1 represent the approximate

amounts of 140 used in the experiment and the dry weight of the

32

leaf tissue (air dried for 1 week at 28°). Using BaluCOB with
a Specific activity (S.A.) of 52.6 nc/mg and assuming the leaf
tissue volume negligible in a total chamber volume of 375 ml.
the final 002 concentration of the samples was between 0.2%

and 0.3%.

Table 1
Photosynthesis Experiment with Tobacco Leaves

Sample Time of Pretreatment Amt. of 170 Dry Wt.

 

 

No. PS , figcurie mg.
l 4" 1 hr. water 328 384
2 4" 1 hr. OH—PMSulfonate 197 391
3 ll" 1 hr. water 180 413
4 11” 1 hr. OH-PMSulfonate 160 471
5 30“ 1 hr. water 234 431
6 30" 1 hr. OH-PMSulfonate 178 364
7 60" 1 hr, water 470 378
8 60" 1 hr, OHPPMSulfonate 282 388

Photogynthesis Egpppiments with Algae:

The normal photosynthetic experiments with algae were
similar to those of Bassham and Calvin (9). The cells, in a
log phase of growth. were centrifuged from the medium at
1000 x g for 10 minutes. The pellet was resuSpended in a
minimum of distilled water and transferred to a pointed centri-
fuge tube and again centrifuged 10 min at 1000 x g. The packed
cell volume was recorded and then the cells were resuSpended
in the reaction medium (usually 0.001 M phOSphate at pH 6.5) so
that a final suSpension of 1% algae cells (v/v) was obtained.

33

The time-rate experiments were performed in a small
"lollipop" containing 15-20 ml of suspended algal cells. White
light of 3000 ft-c was obtained from two 300 watt Kenrad
reflector floodlamps. positioned perpendicular to the plane of
the lollipOp. one on each side. The lollipOp was suSpended in
a water bath maintained at 20°. Five-liter diphtheria toxin
culture bottles, filled with distilled water. were placed bee
tween the lamps and the lollipOp in order to absorb the heat
from the lamps. Air from the compressed air line. passing
through a cotton filter, gently aerated the cultures during
the five minute preincubation period. At zero time 100 pl of
HluCOB' in 0.1 N KOH (0.5 uc/ml) was introduced into the
suspension and the lollipop briskly shaken in the light path.

At various time intervals, approximately 2-4 ml aliquots
were drained into hot absolute methanol so that the final
concentration was about 70-80% methanol. After gentle
boiling. acidification with acetic acid, and aeration with non-
radioactive 002. a 0.1 or 0.2 m1 aliquot of the extracts was
counted in 15 ml of scintillation fluid suitable for aqueous
solutions (51). Total fixation was calculated as counts per
minute per ml of algal suspension.

When photosynthesis experiments in colored light were
performed. several modifications of the above procedure produced
excellent results. The algae suspension in the lollipop was

0.5% rather than 1.0% since the light intensities of the

34

filtered light sources were quite low. Filter systems and

light sources identical to those described for the culture
conditions provided the incident radiation for these experiments
(Figure 2). The distance between the light source and the face
of the lollipOp determined the light intensity which was
approximately measured with a Weston photometer.

The lollipOp was illuminated from only one side and the
experiments were performed in total darkness except for the
filtered light used in the experiment. A gentle stream of cold
air maintained the temperature of the lollipop at approxi-
mately 20° during long experiments (10 min). All other pro-
cedures for handling the algae. isotOpes. and killed samples,

were followed as described above.

Separation and Identification of Photosynthetic Products:

The two-dimensional paper chromatographic techniques of
Benson et a1. (12) routinely were used for the photosynthetic
fingerprint of the percent distribution of 1“0 among the
products. A suitable aliquot of the methanolawater extract was
evaporated to a very small volume (under reduced pressure at
30a35°). This was Spotted on Whatman #1 paper for chroma-
tography and the chromatogram developed in the first direction
with water-saturated phenol and in the second direction with
butanol-propionic acid-water; After drying the approximate
glycolate area on the chromatograms was sprayed with 0.1 M

NaH003 to prevent glycolate sublimation. EXposure of the dried

35

chromatogram to Kodak NO-Screen X-ray film for 2%-3 weeks
revealed the location of the radioactive Spots which were then
counted with a thin window (DuPont Mylar film) gas flow
counter using a Nuclear Chicago scaler (Model 161A). Helium,
passing through ethanol at 0° provided the gas for the
counting chamber. Since only the relative distribution of the
radioactivity on a chromatogram was needed, the absolute
efficiency of the system was never accurately determined;
however. an approximate value of 10% efficiency was used for
gross approximations. Cochromatography of radioactive Spots
with authentic samples and the use of Spray reagents permitted
positive identification of unknown compounds.

When larger samples of a particular compound were needed
for degradation procedures, ion-exchange resins provided the
easiest initial purification. The technique of Plaisted (86),
using Dowex-SO-HI. effectively separated the amino acids from
the other plant material. Further purification was achieved
using two-dimensional chromatography on Whatman #3 with the
solvents described above. Water eluted the purified Spots from
the chromatogram into test tubes.

Zelitch's procedure (139) with Dowex-l-acetate columns was
employed to separate glycolate and phosphoglycerate. 0f the
fractions collected. one contained glycolate and the other
phOSphoglycerate (PGA). These fractions were further purified
by paper chromatography in a basic solvent system (butanol: 95%

36

ethanol:water:diethylamine. 80:10:20:1). The chromatography
removed from the radioactive acids small quantities of unlabeled

plant substances also present in these fractions.

Colorimetric Assays:

The colorimetric assay of Calkins (18) with 2.7mdihydroxyu
naphthalene produced a chromophore with glycolic acid which
absorbs at 540 mp. In the assay 0.2 ml of sample was boiled
for 20 min with 2 ml of a 0.01% solution of the naphthalenediol
in concentrated sulfuric acid. The color reSponse was linear
from 0 to 0.1 pmole glycolate. The sample was diluted with 4
ml of 2 N sulfuric acid and the absorbance of the solution
measured with the Coleman Jr. Spectrophotometer.

Hydrolysis of 3-phosphoglycerate was necessary so that the
free glycerate could be degraded and assayed colorimetrically.
Fractions containing PGA, or the Spots eluted from chromatograms
were combined and taken to dryness. The residue was dissolved
in a minimum volume of water and titrated to the bluengreen
endpoint of bromthymol blue indicator (approximately pH 7.5«
8.0). A 0.1 ml aliquot of neutralized alkaline phosphatase
solution (2-3 mg per 10 ml) completely hydrolyzed the PGA in 2
hr at 30°. The free glycerate was then purified by paper
chromatography in the basic solvent system described above.

The procedure. outlined by Bartlett (7). for a quantitative
glycerate determination with chromotrOpic acid (4,5-dihydroxy-

2.7-naphthalenedisulfonic acid) produced a good reproducable

37

color with a maximum absorbance at 690 mu. A 0.2 ml sample
was heated in a boiling water bath for 30 min with 5.8 ml of
the chromotrOpic acid solution (0.01 % chromotrOpic acid in
concentrated sulfuric acid). After cooling the colored
solution gave a linear absorbancy reSponse with the Coleman

Jr. SpectrOphotometer from 0.01-0.15 pmoles Of free glycerate.

Degradation Procedures:
The modified degradation technique of Sakami. described

by Chang and Talbert (20) was equally effective for degrading
serine and free glycerate. The degradation vessel was a 50 ml,
3 necked, peareshaped flask fitted with a separatory funnel for
adding reagents. a condenser and attached C02 trap, and an inlet
tube for aeration. The degradation followed three separate

steps outlined in the following procedure.

Step one: Cleavage of serine into one mole each of 002. formate.
and formaldehyde.

To the flask was added radioactive glycerate or serine.
0.2 mmole cold carrier (21 mg serine), and 2 ml 0.5 M phOSphate
at pH 5.8. Then, to the closed system, 107 mg NaIOg in 3 ml
water was introduced and the system was aerated 1 hr at room
temperature. Most of the excess periodate was consumed by
reaction with an additional 0.2 mmole of carrier during another

hour of aeration.

38

Step two: Oxidation of formate to C02 (carbon 2 of the acid).

To the flask was added 1.5 ml 3.0 M phosphate at pH 2.5.
Then one gram of HgClz, dissolved in 5 ml hot water, was
introduced into the closed system through the funnel. The system

was aerated for 1 hr while boiling gently.

Step three: Oxidation Of formaldehyde to C02 (carbon 3 of the
acid).

To the flask was added 1 ml 5% AgNO3; the system was
closed. and 1.5 g K23208 in 5 ml water was added through the
funnel. First the solution was aerated for 45 min with gentle
heating. then aeration continued 30 min longer while boiling.

After each step the 5 ml of the COzntrapping solution (128)
(27 ml redistilled phenylethylamine. 27 m1 absolute methanol.
500 mg POP, and 10 mg POPOP taken to a final volume of 100 ml
with toluene) was quantitatively transferred to a polyethylene
counting vial. The trap was washed with scintillation fluid
(5 g POP and 100 mg POPOP per liter of toluene) three times so
that the final volume in the counting vial approximated 15 ml.

The degradation of glycine followed the exact procedure of
Chang and Talbert (20). Ninhydrin released the carboxyl carbon
of glycine at room temperature, and boiling with potassium
persulfate in the presence of AgN03 oxidized the second carbon
to C02. The same degradation apparatus and 002 trapping system

were used as described for the serine degradation.

39

The degradation of glycolate proceeded according to the
technique outlined by Zelitch (139) with slight modifications.
The degradation was performed in the degradation vessel
previously described rather than a Warburg flask. The trap
contained 5 m1 of the amine COz-trapping solution rather than
KOH as used by Zelitch. This degradation also proceeded in

three steps:

Step one: Oxidation of glycolate to glyoxylate.

To the flask were added 1.0 ml 0.1 M phosphate at pH 8.0.
1.0 ml 6 x 10‘” M FMN. and approximately 2,000-5.000 cpm of
the radioactive glycolate. Gentle aeration with 02 was started
and 0.2 m1 of a glycolate oxidase preparation was added through
the funnel with a total volume of 2 ml water. After 4.0 ml
sodium glycolate carrier solution (500 ug/ml) were added.
aeration was continued for 1.5 hr. The oxidase prepared from
tobacco sap. was the ammonium sulfate fraction precipitated be»

tween 15-30 g (NH4)2804 per 100 ml solution.

Step two: Cleavage of glyoxylate into one mole each of 002
and formate.

Through the funnel was added 0.6 ml of 0.2 N (NHh)4Ce(SOu)4
in 2 N sulfuric acid, and the solution boiled gently 1 hr
without heating. Thus, the possible distillation of formic acid
at this pH was minimized. The flask was cooled to room temperae
ture and aerated gently with air for 30 min to quantitatively

remove the liberated 002. Care must be taken to avoid excessive

4O

bubbling. since the protein solution may foam through the

condenser and ruin the COz-trap.

Step three: Oxidation of the formate to 002 (carbon 2 of

 

glycine)

After addition Of a second COZ-trap. 2 g HgClz in 10 ml
hot water was added through the funnel. This mixture was
boiled gently for 1 hr with continuous aeration.

For each degradation the counting efficiency with the
amine cog-trapping solution approximated 55-60% as compared with
20% with the Cab-O-Sil gel counting of KOH solutions. The
distinct advantage of being able to count the entire amount
of 002 trapped, rather than a small aliquot of XOR solution
allowed the use of fewer counts per degradation.

The calculation of percent yield was Obtained by counting
an aliquot of the non-degraded material and comparing it with
the total recovered. The percent distribution of 140 within
the compound was directly calculated from the amounts of 190
recovered at each degradation step. Results of standard
<1esradations with acids of known radioactive distribution
(Tables 2 and 3) indicated the reliability of the glycerate.
glycine, and serine degradations. In all cases at least two
desradations were run on each standard sample. but more were run
(”1 Slycolate samples to confirm the low recovery of the second

carbon atom .

41

 

 

 

 

 

Table 2
Percent Distribution of 140 in Standard 0. Compounds

Percent_;uC

Compound Percent Recovery Cl 02 03
3-PGA-U-1uc 1 95 33.9 33.3 32.7
2 96 34.7 33.1 31.4
Serine-1-1”C 1 97 92.6 1.6 5.8
2 -- 95.3 1.0 3.3
Serineu3-1uC 1 96 0.4 6.8 92.8
2 -" 009 1117 9701+

 

 

 

The total yield of 170 from glycolate-Z-luc was consistently
low and varied between 90-97 %. The recovery of the first
carbon of glycolate was excellent (Table 3). It was concluded
that the most valid calculation for the 170 distribution in

f 11+C recovered in

glycolate would be to determine the percent a
the first carbon with reSpect to the total activity added to the
reaction rather than with respect to the total 140 recovered.
Since the total recovered was always less than 100%, due to poor
yields of the second carbon. calculations on the amount of
recovered radioactivity would increase the percent of 14C in the
first carbon of glycolate. The effect of this argument is
presented in Table 3. Uniformly labeled glycolate yielded a
55/45 (01/02) ratio if the distribution was based on total 14C

recovered in the degradation. This ratio became 51/49 if the

42

distribution in Cl was based on the total 140 added to the

reaction and the C2 was calculated by difference.

Percent Distribution

Table 3

of 140 in Standard C2 Compounds

 

Compound
Glycine-l—
G1 1. -2-
yclfic
Glycplate-
Glycplate-

Glycolite-

[.1

NH NH

 

Percent

Recovery

100
100

96

100

94.4

90.9
97.3

94.6

Percent of 140

 

 

Recovered

C1 02
87.4 12.4
84.8 15.1
1.3 98.7
99.5 0.5
99.0 1.0
13.8 86.2
14.4 85.6
54 5 45.5

 

Percent of 140

Added

 

Cl

99.5
93.5

12. 6
14.0

51.4

C2

 

Calculation of Sppcific Activities
From the concentration of the acid solutions. determined
colorimetrically. as pmoleS/ml, and the activity of the solution
evaluated as counts per minute per ml (Cpm/ml) of the same
solution, the specific activity as cpmflpmole acid was determined
by division.
pmole C) resulted as the product of the total specific activity

and the percent of the 1”C in a particular carbon. A convenient

for Glycerate and Glycolate:

The Specific activity per pmole of carbon (apm/

check on this calculation was the sum of all the Specific

43

activities of the carbons in a compound, which must equal the

specific activity of the whole compound.

Sonication Of Algae:

The algae were harvested as previously described for the
photosynthesis experiments. The packed cell volume was diluted
to a 30% (v/v) suSpension in a designated medium. The sonic
converter (Branson Sonifier, Heat Systems (Melville. L.I.)) was
a probe instrument and rupture of the cells was accomplished
in "rosette cells" supplied by Heat Systems Co. Routinely 15
ml of algal suspension at pH 7.5-8.5 in the 25 ml rosette cell
was cooled in an ethanol-water-ice bath at -5°. The regular tip
of the sonicator was positioned as close to the bottom of the
cell as possible, and the instrument Operated at the maximum
output level so that the ammeter indicated 0.55~O.65 amps. The
Sonifier ruptured 90% of the Chlamydomonas cells after five
minutes. but only 60-70 % of the Chlorella or Ankistrodesmus
cells after 10-15 min. The apparent turbidity of the suspension
decreased as the cells were broken and particle size decreased.
The pH values of the broken suspensions varied as indicated by

the data of Table 4.

Table 4

pH of Algal Sus ensions Before and After Sonication

 

SuspendinggMedium pH Before Sonication pH After Sonication

 

Water 6.5 5.7-5.8

0.06 M phOSphate
buffer 8.3 7.5

0.02 M cacodylate
buffer 6.3 6.3

 

 

 

It was observed that at pH values less than pH 5.5. the sus-
pension would sometimes coagulate and greatly reduce the
efficiency of the sonication.

Centrifugation at 3.5 x 10“

x g for 10 min removed the un-
broken cells and cellular debris. The clear dark green super-
nate was then used in experiments as a cell extract or

fractionated for a Specific enzymatic activity.

Enzyme Assanyrocedures:

 

Glycolate Oxidase:
The manometric technique (21) employed 0.5 ml of 0.1 M

phosphate buffer at pH 8.3. 0.5 ml 6 x 10““

M FMN, 0.5 ml water.
and 1 ml cell extract (pH 7.5-8.0) in the main portion of the
Warburg vessel. The side arm contained 0.5 ml 2 x 10""2 M Na-
glycolate and was added with the main chamber at zero time.

The oxidase activity was evaluated from the volume of 02 uptake

per unit time.

44

731“

185'

45

In the very sensitive isotopic technique a known amount
of glycolate-z-luc was added to the crude cell sonicate (not
centrifuged) at zero time. The complete mixture was incubated
in the light with gentle shaking and aliquots killed by mixing
them with hat methanol. Glycolate oxidase activity was
measured by the identification of glycine and serine on two
dimensional chromatography Of this reaction mixture. The
relative amounts of glycine and serine were compared with the

amount of glycolate added as substrate.

Phasphoglzcolate_phOSphatase:
The phosphoglycolate phosphatase assay was that described

by In et al. (131). except that cacodylate rather than Tris-
acetate buffer was used. The mixture contained 0:1 ml buffer
(0.1 s cacodylate plus 0.02 s hgsog at pH 6.3); 1.0 ml 0.01 n
phosphoglycolate and 1.0 ml enzyme. The mixture was incubated
at 30° for 30 min after the addition of the enzyme, and then A
was killed with 1.0 ml of TCA (10%). After centrifugation. 1
ml of the supernate was assayed for orthophOSphate (25).

Spectrophotometric Assags:

The Gilford automatic cuvette changer and absorbance con-
verter. Beckman DU spectrophotometer. and Sargent recorder were

used to measure the rates of the following enzymatic reactions.

46

Glycolate Oxidage:

A spectrophotometric assay observed the increased
absorbancy at 324 mp due to the formation of glyoxylate phenyl-
hydrazone. This reaction mixture contained: 2 ml 0.1 M
phosphate pH 8.3: 0.1 ml 0.1 M phenylhydrazine hydrochloride
adjusted to pH 6.8: 0.1 ml 0.1 M.cysteine: 0.5 ml enzyme (cell
extract): 0.3 ml 0.1 M glycolate (total volume of assay = 3 ml).
The increased absorbancy formed during a given time interval

was defined as glycolate oxidase activity.

Glzogzlate Reductase: .
The assay procedure of Zelitch (135) followed the oxida-

tion of reduced pyridine_nucleotide2 measured as an absorbancy
reduction at 340 up after the addition Of glyoxylate. The
assay mixture contained: 1 ml 0.1 M phOSphate pH 6.5. 0.5 ml
0.1 M glyoxylate, 0.05 ml 0.02 M NADR. 0.1 ml cell extract, and
water to 3 m1.

Isocitrate Depydrogenase:
Isocitrate dehydrogenase actiVity was detected using the

assay procedure described by Syrettpgt‘gl‘ (102). The assay
followed the rate of NADP reduction measured as increasing
absorbancy at 340 mp. The assay mixture contained: 1 ml 0.1 M
phosphate pH 7.5; 1 ml 0.05 M MgClz. 0.1 m1 0;06 M cysteine.

u)
D

1')

O‘

A)

47

0.05 ml 0.02 M NADP, 0.2 ml enzyme preparation (cell extract),
and 0.1 ml 0.1 M isocitrate used to start the reaction.

Extraction and Determination of Chlorophyll:

The 80% acetone chlorOphyll extraction which has been used
for extracting chlorophyll from chloroplast preparations (2)
did not extract much of the chlorOphyll from algal extracts.
The extinction coefficients. which Mackinney (64) has reported
for absolute methanol solutions Of Chlorophyll, made it possible
to determine the chlorophyll concentration directly on a
methanol extract from algae. Ten ml of a 0.5 % algal suSpension
provided sufficient chlorophyll for the SpectrOphOtometric
determination. The algae were centrifuged in an International
Clinical centrifuge at maximum Speed for 3 min. Immediately
the supernate was discarded and the tube inverted on an
absorbant surface so that the excess water would drain from the
sides of the tube, After several minutes. the cells were rem
suspended in 3 ml of absolute methanol (using a Vortex mixer),
and were placed in stOppered centrifuge tubes in the dark for at
least 2 hr to insure complete extraction of the pigment. After
centrifugation, absorbancy measurements on the green supernate
were made with the Beckman DU SpectrOphotometer. From
Mackinney's extinction coefficients at 665 mp and 650 mp the

following equations for chlorOphyll concentration were calculated:

48

mg Chlorophyll a per liter = 16.2 Abs665 - 7.53 Ab3650

mg ChlorOphyll p per liter = 30.6 Abséso - 11.3 Abs665
The absorbancy measurements at these wavelengths were converted
to mg chlorophyll/liter, and a final value as mg chlorOphyll/
ml of packed cells was obtained by multiplying by 0.06 to
account for the 3 ml of methanol used and the volume of packed

cells (0.05 ml).

Measurement of in vivo Absopption Spectra:

Light scattering presents the main problem in the accurate
measurement of the Spectral absorption of pp zggp algal sus-
pensions. The technique Of Shibata,et_al$ (96) employed a
neutral density filter made of filter paper saturated with
mineral oil which was placed between the light source and the
cuvette. It was found that a double thickness of waxed
weighing paper (Schleicher and Schuell CO. NO. B-2) placed
next to the cuvette on the face toward the light source most
effectively elucidated the fine structure of the pp 3319
chlorophyll spectrum. The ratio of chlorOphyll a/b absorption
could be visibly evaluated from Spectra measured on the Cary
15 spectrophotometer. Good resolution was achieved with a
slit width of approximately 0.7 mm. Under these conditions
the Absorbance of a l % algal suSpension was approximately 0.5

Absorbancy units at 750 mu.

5
'OC'.
-vvu

\f’fl‘
w‘uv:
0

“7"“ a 1

NV I

49

Growth rates of algal suspensions were obtained by
measuring aliquots of algal cultures at various time intervals
after innoculation. The log of the Absorbancy of the sus-
pension measured at 680 mu with the Coleman Jr. Spectro-
photometer. was plotted against time. Typical data revealed

whether a culture was in a log phase or a lag phase of growth.

r... u u
t I
2. Av
« Ch
a.

d

1: e
ailat

. 1
-919

“ fi'p
hék

x.“

1'“

RESULTS AND DISCUSSION

 

The Rate of Glycolate Formation in Tobacco:

In the photosynthetic experiments with tobacco leaves, as
described in the methods section, the linear rate of 14002
fixation observed during the first 60 sec in the control experi-
ment (Figure 3) indicates that the tissues did not run out of
14002. i.e. 14002 did not become rate-limiting. It is
important for the arguments about the movement of 140 into the
metabolic pools, as represented by glycolate, that these leaves
were at all times saturated with 17002. The abnormally high rate
of fixation by the inhibited tissue for the 30 second experiment
might have been due to a difference in the biological activity
of the tissue rather than its Size, since as seen in Table l,
the dry weight of the tissue was not larger than the other
samples. That the inhibitor was taken up by this particular
sample was evident in the product distribution.

, The percent distribution of 140 fixed into the various
products of photosynthesis from 4 to 60 seconds are shown in
Figure 4. The percent of 140 fixed into glycolate and glycerate
increased with time, but that fixed into PGA decreased. These
data agree with the classical results obtained by Bassham and“
Calvin (9), indicating that initially PGA contains most of the

newly fixed 1“’0 and that other compounds are formed subsequently.

50

Figpre 3
Total Fixation of 14002 by Tobacco Leaves

 

Leaves photosynthesized 14002 in full sun-
light.

Closed symbols: Control (1 hr pretreatment
with water)

Open symbols: Inhibitor (1 hr pretreatment
with 0.01 M OH-PMSulfonate

51

 

(+) OH-PMSulfonate

 

Control

 

Figure 3

 

Tobacco Leaves

60

The

 

 

200_.
150..
1OO_5

Amlo_ x B\ov aoapomam mansaom ampaziaoamnpms OH voxfim owF Hopes

Time (sec)

52

Achesocaamzaumo
a Ho.o spas paeaaeoancaa as av nonapassH .nHopaan coco

Ahead: new: psospaoauona H: av Honpaoo anopaaw domoao

.pstHSSm HHsm ma manna Oomdmozpaamoposa moaned

 

mobmoq ooompoa an Oasnom
opaHooaHo dam .opaaoomau .<om ad 03H mo soapdnaauman paOOHom

a cammHm

53

noowv mafia

 

 

   

opscomadmzmlmoA+v

Wewqoowflw

 

omeOMHszmumoA+v

   
    

Hopp:oo

MadeOMHw

 

  

HOMPCO O

oemnomaamzmlmoa+v

ma<mmOwnwomMmomm

    

0—

10m

 

 

e onswdm

uOIlOBIJ qunIos madam-Iousqiam u: pext; otl quoaxag
54

55

These data also demonstrate that the inhibitor. OH—PMSulfonate,
effectively blocked glycolate oxidase as evidenced by the
gigantic accumulation of 11‘1C in this acid. The demonstration
that movement of 140 into the rest of the glycolate pathway was
also blocked by OHAPMSulfonate is shown in Figure 5. A re-
ciprocal relationship existed between the accumulation of
glycolate-140 in the presence of the inhibitor and the decrease
in the 1“(C within the products of the glycolate pathway. glycine,
serine. and sucrose. I conclude. therefore, from 140 pool
sizes that the inhibitor. OH-PMSulfonate. as described by
Zelitch (134). blocks glycolate oxidase with the resultant
accumulation of 140 in glycolate. Further. I confirm that PGA
and not glycolate is the initial product of photosynthetic 002
fixation. even in the presence of an inhibitor which favors
glycolate accumulation.

In order to more completely defend this latter conclusion.
it was necessary also to evaluate the relative Specific
activities of PGA. glycolate. and glycerate. The relative
total amounts of radioactivity isolated in these compounds
(Figure 6) suggest that not only did the inhibitor cause the
accumulation of glycolate but it also reduced the amount of free
glycerate. The latter conclusion is to be expected if
glycerate were a product Of the glycolate pathway. although
labeling data to be presented does not support this argument.

Figpre 5

Percent Distribution of 11+C in Glycolate. Glycine. Serine.
and Sucrose Formed by Tobacco Leaves

Leaves photosynthesized 14002 in full sunlight.
Closed symbols: Control (1 hr pretreatment with water)

Open symbols: Inhibitor (1 hr pretreatment with 0.01 M
OH—PMSulfonate)

56

Figure 5

 

50

5'
.L

J

U
10

(D
O

l

1

Percent 140 fixed in methanol-water soluble fraction
c>
1 l

 

GLYCOLATE GLYCINE,

SUCROSE

(+) OH-PMSulfonate

 

Control

W

l l l

 

SERINE,

Control

(+) OHmPMSulfonate

      

 

1 1 l
4 11 30 6O 4 11

Time (sec)

57

30

6O

 

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.pmmfiasam Adam aa douamoSCSAmopoaa mobaoq

 

needed
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b camwaa

58

 

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60

That most of the free glycerate did not arise from the hydrolysis
of PGA during the killing procedure is indicated by the changes
in the ratios of PGA/glycerate (Figure 6). The ratio of PGA/
glycerate was not approximately constant as would be expected
from phOSphatase hydrolysis of PGA during killing.

The Specific activities of the Chromatographically pure
samples of PGA, glycerate, and glycolate. are presented in
Table 5. These values can be corrected to approximate disinte-
grations per minute by multiplying by 2. since we observed 50-
60% counting efficiency in our scintillation counting procedure.
These data, as plotted in Figure 7. demonstrate that the S.A. of
PGA immediately reached a plateau (within 4 seconds) and that by
30 or 60 seconds even the S.A. of glycerate and glycolate were
approximately that of PGA. These data do not confirm Zelitch's
Claim (139) that the S.A. of glycolate is greater than the S.A.
of PGA. Zelitch used experimental times of 2 and 5 minutes and
killing times of 30 seconds in 12002 air.

Since the Specific activity of the carboxyl group of PGA
and the carbon atoms of glycolate must be compared. both com-
pounds and also glycerate were degraded as described in the
methods section. The results (Tables 6 and 7) indicate that PGA
and glycerate were predominantly carboxyl labeled at short
times, but that they became almost uniformly labeled by 60

Seconds. The glycolate (Table 8), however. showed at all times

 

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moa N o.m moa N 5.5 moa N o.m opmsomasmzmimo as
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w OHDRB

61

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62

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63

Percent Distribution of 140 in 3-PhOSphoglycerate
Isolated From Tobacco Leaves After Photosynthesis in 1

Table 6

4002

 

Time of PS

Pretreatment

 

 

4w
4"

11"
ll"

30"
30"

60"
60"

Water

OH-PMSulfonate

Water

OH-PMSulfonate

Water

OHHPMSulfonate

Water

OH-PMSulfanate

% Distribution of 140

Cl
83.2
87.4

64.9
64.0

51.1
49.6

36.7
43.1

C2

809
7.1

16.1
16.5

23.0
28.1

33.3
28.8

03
7.9
3.5

19.0
19.5

25.9
22.3

30.1
28.1

 

64

Table 7

Percent Distribution of lac in Glycerate
Isolated From Tobacco Leaves After Photosynthesis in 11+C02

 

% Distribution of 140

 

 

Time of PS Pretreatment C1 02 03
4" Water 83.9 9.2 6.9
4" OH-PMSulfonate 89.1 4.9 6.0
11" Water 73.5 12.5* 13.0*
11" OH-PMSulfonate 72.6 15.0 12.4
30" Water 54.1 27.4 18.5
30" OHAPMSulfonate 42.9 29.9 26.2
60" Water 34.4 35.9 29.7
60" OH—PMSulfonate 40.5 34.0 25.6

 

% error for these averages was generally 5% although those
marked * were 10%; the values were averaged from two separate
degradations.

65

 

66

 

 

 

 

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cadence Sana cepmmomH opaHOOhHw SH OSH mo SOHpannpmHQ paooaom

m eHoea

67

equal distribution of the label in both carbon atoms. Using the
S.A. data in Table 5 and the percent distribution data in Tables
6, 7, and 8. the S.A. of the individual carbon atoms of these
compounds have been calculated (Table 9). Thus, the S.A.. 5.7 x
107 cpm/pmole C, for the carboxyl carbon of PGA is 100 times
greater than the S.A.. 6.0 x 105 cpm/pmole C, of either carbon of
glycolate at 4 seconds. The rapidity of photosynthesis in this
tissue is emphasized by observing that the S.A. of these same
carbons was almost equal after 60 seconds. The very high initial
S.A. of PGA confirms that carboxy labeled PGA was formed
initially by the carboxylation reaction of the photosynthetic
carbon cycle. Labeling of glycolate occurred subsequently. That
Zelitch observed a S.A. of glycolate higher than PGA must reflect
a rapid incorporation of non-radioactive 002 into his samples
during his long killing procedure. thus reducing first the
Specific activity of PGA.

The effect of the hydroxysulfonate on glycerate formation
invites Speculation. AS already mentioned the inhibition of
glycerate formation by OHHPMSulfonate (Figures 4 and 6) might be
explained as a result of blocking the glycolate pathway. However.
if the glycolate pathway were contributing to the pool of free
glycerate. the distribution of label in glycerate Should be
either uniform or more quickly randomized in the control leaves
than when the inhibitor was present. This was not observed

(Tables 6 and 7). The lower S.A. Of glycerate in the presence

Table 9

Specific Activities of Carbon Atoms in PGA, Glycerate. and Glyco-
late Which Were Isolated From TpRacco Leaves After Photosynthesis

 

 

 

 

 

 

 

 

 

 

 

 

in 002
Time of PS Pretreatment Specific Activity (cpm/umole C)
3-phosphoglycerate
Cl 02 C3
4" Water 5.7 x 107 6.1 x 106 5.4 x 102
4" OH-PMSulfonate 2.6 x 107 2.1 x 106 1.1 x 10
11" Water 1.2 x 107 3.1 x 102 3.6 x 106
ll" OH-PMSulfonate 1.3 x 107 3.3 x 10 3.9 x 106
30" Water 7.7 x 10? 3.5 x 103 3.9 x 102
30" OH—PMSulfonate 5.5 x 10 3.1 x 10 2.5 x 10
60" Water 2.9 x 107 2.7 x 107 2.5 x 107
60" OH—PMSulfonate 3.6 x 107 2.4 x 107 2.4 x 107
Glycerate
C1 C2 C3 .
4'1 Water 6.1 x 105 6.7 x 10: 5.0 x 10:
4" OH-PMSulfonate 2.7 x 105 1.5 x 10 1.8 x 10
11" Water 8.1 x 105 1.4 x 10 1.5.x log
11" OH—PMSulfonate 1.8 x 105 3.8 x 10 3.1 x 10
30" Water 5.3 x 102 2.7 x 102 1.8 x 102
30" OH-PMSulfonate 1.7 x 10 1.1 x 10 1.0 x 10
60" Water 7.6 x 106 7.9 x 102 6.6 x 102
60" OH-PMSulfonate 2.5 x 106 2.2 x 10 1.6 x 10
Glycolate
4" Water 6.0 x 105 7.9 x 105
4*1 OH-PMSulfonate 4.0 x 105 3.7 x 105
11" Water 1.4 x 106 1.3 x 102
ll" OH-PMSulfonate 1.2 x 106 1.2 x lo
30" Water 6.7 x 102 6.3 x 102
30" OH-PMSulfonate 8.3 x 10 7.7 x 10
60" Water 8.4 x 106 7.7 x 106
60" OH-PMSulfonate 1.6 x 107 1.7 x 107

68

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69

of OH—PMSulfonate (Table 5) suggests that the inhibitor decreased
the amount of 1&002 incorporation into glycerate. This block in
glycerate formation may not have been solely in the glycolate
pathway, since the lL’Cndistribution in the remaining glycerate
does not correspond to that expected from glycolate. If the
radioactivity, passing through the glycolate pathway, did not
contribute significantly to the pool of glycerate until after 60
seconds, then one may not expect to see more rapid randomization
of glycerate in the control system than in the inhibited one. A
possible solution to this problem might be found in the degradaa
tion of serine from these experiments. Rabson, Tolbert, and
Kearny (90) have shown uniform labeling of serine after 20 sec by
the intact leaf. One should be able to differentiate between
serine from the glycolate pathway in the control experiments and
serine formed from glycerate in the inhibited system. Similarly
Chang and Tolbert (20) were able to differentiate between two
reservoirs of serine, one in the chlor0p1asts and one in the

cytOplasm.

Engymes in Algae:
Glycolate Oxidase:

For many reasons, investigators have tacitly assumed that
the glycolate pathway Operated in green algae. The ubiquitous
distribution of glycolate oxidase in higher plants has been well
demonstrated. Algae also rapidly label glycolate, glycine, and

serine during photosynthesis as do higher plants. However,

BEE

70

there have been no literature citations to this enzyme from
algae except for three preliminary reports (40.53.78) which
suggest that if the oxidase is present it is likely different
from that of higher plants. Therefore experiments were run to
attempt to isolate and characterize glycolate oxidase from
sonicated preparations of Chlorella, Chlamydomonas, and
Ankistrodesmus.

Extracts from cells of 2 or 3 day old cultures were pre-
pared as described in the methods section. Before sonication
the cells were suspended in either, 0.1 M phosphate buffer at
pH 8.3, 0.1 M phosphate at pH 8.3 containing 2 x 10-2 Na-
glycolate, or 0.1 M phosphate at pH 8.3 containing 2 x 10‘”2
cysteine. After sonication, these suSpensions were centrifuged
so that the original crude extract, the supernatant fluid and
the residue could be assayed manometrically. No glycolate
oxidase activity was observed in any of these preparations from
any of the algal strains under any conditions. One-half ml of
an active glycolate oxidase preparation from tobacco when added
to flasks which contained samples of the above extracts
demonstrated no inhibition of activity. Addition of glycolate
oxidase from tobacco to the cell suSpension before sonication
indicated a 10% reduction of the initial activity (detected
manometrically) after the cells were sonicated. No oxidase .
activity was observed as glycolate dependent 02 uptake. Flushing
the flasks with pure 02 gas for 5 minutes before the eXperiment

also did not promote any activity.

71

When glycolate-Z-luc was added to these algal extracts,
little or none of the isotOpe was converted to glycine or serine
or any other compound as measured by paper chromatography.

These results are in agreement with a recent report of
Whittingham (125). To test the possibility that the transaminase,
converting glyoxylate to glycine, might have been inactivated,
and that glyoxylate cannot be easily detected by paper chroma-
tography, I used an assay for glycolate oxidase which follows

the rate of glyoxylate formation as the accumulation of the
glyoxylate-phenylhydrazone. This assay also did not indicate

any glycolate metabolism by any of the algal extracts. These
experiments were also done with Chlamydomonas cultures at various
stages of growth, lag phase, log phase, and senescent algae, but
no oxidase could be detected in any of the preparations.

In the above experiments numerous controls were run and
they are summarized as follows: (a) No inhibitor for the normal
glycolate oxidase from tobacco leaves was present in the algae
cell extracts. (b) No enzyme activity was detected when the
algal extracts were prepared in the presence of 2 x 10""2 M
glycolate which Baker and Tolbert (6) had used to isolate a form
of glycolate oxidase from etiolated wheat tissue. (0) Oxidase
from tobacco leaves was not significantly destroyed by sonication.
(d) Increasing the partial pressure of oxygen had no effect on
the endogenous respiration. Glycolate oxidase from green leaves
has a low affinity for oxygen and is dependent upon ample oxygen

in the atmosphere (106).

72

Thus, the assumption that the typical glycolate pathway is
Operating in algae must be modified. Certainly the conclusions
from these studies on the oxidase eliminated the possibility that
glycolate metabolism, if it occurs at all in algae, proceeds via
a typical glycolate oxidase. Recently other reports have
suggested that glycolate oxidase in algae may be different from
that in the higher plant (53,78). The theory that glycolate is
excreted by algae, as if it were an end metabolic product (125),

is confirmed by these enzymatic studies reported here.

Isocitrate Dehydrogenase:

 

To evaluate whether or not the lack of glycolate oxidase
activity reflects a general inactivation of algal enzymes during
preparation of extracts, isocitrate dehydrogenase was assayed as
described in the methods. The algal extracts demonstrated a rate
of NADP reduction (30 pmoles NADPH formed per 10 min per 0.1 ml
extract) comparable to that reported by Syrett,et_al; for
Chlorella vulgaris (102). This activity was dependent only on
isocitrate and Specific for NADP, since NAD was completely
inactive. That isocitrate dehydrogenase and other enzymes to be
mentioned below could be detected, under conditions identical
with those used for preparation of glycolate oxidase, demonstrated
that active enzymes were present in the sonicated cell-free

extracts.

73

Glyoxylate Beductase:

 

An assay for glyoxylate reductase also showed the presence
of an active enzyme which was dependent upon NADH, but was in-
active with NADPH. A reaction of 30 pmoles NADH oxidized per 5
min per 0.05 ml extract was observed with glyoxylate substrate.
The enzyme preparation was also capable of reducing hydroxy-
pyruvate with NADH. Laudahn (61) has reported that both the
glyoxylate reductase from spinach leaves and D-glycerate dehydro-
genase from tobacco leaves have approximately the same Km values
of 1.2 x 10‘“ for hydroxypyruvate and 1.2 x 10'2 for glyoxylate
(cf. 9.1 x 10"3 for glyoxylate reported by Zelitch, 135). Thus,
the observation of glyoxylate and hydroxypyruvate reduction by
NADH in algal extracts could indicate activity for both
glyoxylate reductase and D-glycerate dehydrogenase, or that one
enzyme was acting on both substrates. Further isolation and
characterization of these reductive enzymes from the algal ex»
tracts are needed before any conclusions can be made about their

function in algal cells.

Phosphoglycolate Phosphatase:

 

Extracts from Chlorella, Chlamydomonas and Ankistrodesmus
all contained active phosphoglycolate phOSphatase. The phospha-
tase, detected in extracts prepared in 0.02 M cacodylate at pH
6.3, gave a ratio of 20:1 with reSpect to hydrolysis of phOSpho-

glycolate and phosphoglycerate. This ratio compared favorably

Table 10

A Summary: Preparation of Cell-Free Extracts From Algae And
Detection of Enzymes in These Extracts.

Procedure:
1. Cells were grown in 1200 ft-c white light with 0.2 %
002 in air.

2. Algae were harvested and resuSpended (30 % v/v) in
water, 0.1 M phosphate buffer, or buffer plus 0.01 M

glycolate, each at pH 7.5.

3. Cells were sonicated with maximum output at 5°, 15
min for Chlorella or Ankistrodesmus and 5 min for

Chlamydomonas.

4. Extracts were assayed by Warburg reSpirometer, iso~

tOpic products and SpectrOphotometric techniques.

 

Results
GLYCOLATE OXIDASE __________ ABSENT
Phosphoglycolate PhOSphatase _____ Present
NADH:Glyoxylate Reductase ______ Present
Isocitrate Dehydrogenase _______ Present

74

75

with the partially purified phosPhoglycolate phOSphatase isolated
from tobacco leaves which gave a phosphoglycolate/phoSphoglycerate
ratio of 3:1 after an acetone fractionation (131).

As summarized in Table 10, the active enzymes, NADP:
glyoxylate reductase, phOSphoglycolate phOSphatase, and isocitrate
dehydrogenase, have been demonstrated in the sonicates of three
strains of algae. In contrast, no glycolate oxidase was observed
with these same algal preparations. Similar negative results
for glycolate oxidase have also been obtained from sonicated

preparations of Scenedesmus obliquus (Gaffron D-3) and

 

Chlorella pyrenoidosa (Warburg).

Distribution of 14C in Products of the Glycolate Pathway Formed
by Algae During Photosynthesis in 1“C02:

 

During initial 14002 fixation by algae, glycolate is
uniformly labeled (107) and PGA is carboxyl labeled (9) as is
also observed in higher plants. However, the absence of glycolate
oxidase (as reported in the previous section) raised speculation
about 14C distribution in glycine and serine. Therefore, the 1”C
distribution in the carbons of phosphoglycerate, glycine and
serine, formed by Chlamydomonas reinhardtii and Chlorella

 

pyrenoidosa (Chick) during short time photosynthesis was determined.
The isolation and degradation procedures for PGA, glycine,
and serine were described in the methods section. Both PGA and

serine were identically carboxyl labeled (Table ll). The formation

HI
- nu
PV

Table 11

Percent Distribution of 140 in the Carbons of PhOSphoglycerate, Glycine
and Serine Formed by Algae During Photosynthesis.

 

A 2 % suspension i4 algae in 0.001 M phOSphate buffer at pH 6.5

 

 

photosynthesized 002 in white light (3000 ft-c).
% Distribution of 140
.31.. 3.2. .91.

Chlorella_pyrenoidosa

( 7 sec PS) P-glycerate 84 4 12
Serine 78 6 16.

(12 sec PS) P-glycerate 82 3 l5.
Serine 80 4 l6
Glycine 50 50

Chlamydomonas reinhardtii

(12 sec PS) P~glycerate 79 4 1?
Serine 69 7 2“
Glycine 49 51

Soybean*

(20 sec PS) P-glycerate 74 ll 16
Serine 28 38 34

*From Babson et a1. (90)

76

77

of carboxyl labeled serine by algae differs markedly from the
formation of uniformly labeled serine by soybean. In both plant
tissues, the PGA was still carboxyl labeled. (See data and
reference cited in Table 11). In these experiments the labeling
of serine in algae did not become more uniform than that of PGA.

These results further support the concept that the glycolate
pathway does not function in algae. As is seen in Figure 1, an
alternate pathway for serine formation via PGA, glycerate, and
hydroxypyruvate, could explain the formation of carboxyl labeled
serine. My observation of an active D-glycerate dehydrogenase W
or glyoxylate reductase in the algal extracts would also be con-
sistent with this hypothesis. Chang and Tolbert (20) have
reported that serine formed by isolated chloroplasts is also
carboxyl labeled; Zak reported that alanine and serine formed by
Chlorella were carboxyl labeled (133).

The glycine formed by the algae was uniformly labeled and
this finding must be compared with uniformly labeled glycolate
and carboxyl labeled serine. Chang and Talbert also observed that
glycine formed by chlor0p1asts was uniformly labeled (20). In
agreement with my results, Zak (133) reported that glycine
formed by Chlorella after 13 sec photosynthesis had 49 1 of its
1“(3 in the 01, while for the same length of time serine contained
82 I, 0 I, and 17 x respectively for C1, C2, C3. This labeling
pattern eliminates the possibility that the reversal of serine

78

hydroxymethylase can account for the glycine production, since the
glycine thus formed would necessarily be carboxyl labeled as is
the serine. Uniformly labeled glycine might come from the
uniformly labeled glycolate, but the absence of glycolate oxidase
and the excretion of glycolate make this possibility unlikely.
As with the work of Whittingham (89,125), the source of glycine
in algae remains unexplained. Perhaps a third, as yet
unrecognized, pathway exists for the production of uniformly
labeled glycine.
Effect ofcx-Hydroxymethanesulfonates On 1["002 Photosynthesis by
Algae:

cx-Hydroxypyridinemethanesulfonate has been used to inhibit
glycolate oxidase and to evaluate the function of the glycolate
pathway in plants (4,134,137,139,l43). Algal experiments were
initiated for a similar purpose. The only report of this in-
hibitor‘s interaction with algae has been from Asadahgt'al; (4)
who observed an inhibition of COZ fixation.

The normal photosynthesis experiments with NaHluC03 were
run with 1.0 % suspensions of Chlamydomonas reinhardtii and
Chlorella pyrenoidosa (Chick) with and without the 0H=PMSulfonate.
Although data with the Chlorella were qualitatively similar, only
data with the Chlamydomonas experiments have been presented
(Tables 3-7 of the Appendix). A comparison of typical radioauto-

graphs from two dimensional chromatography of the methanol-water

79

soluble fraction from tobacco (Figure 8) and Chlamydomonas
(Figure 9) reveals the striking difference between the 1”C dis-
tribution in these two plants when treated with OHnPMSulfonate.
The remainder of this discussion is concerned with a presentation
and interpretation of data obtained from such chromatograms.

The data in Figure 10 indicates a linear in

COZ fixation
rate by the algae for the first 4 minutes of fixation, and that
this rate was greatly stimulated in the presence of 0.001 M OH-
PMSulfonate at pH 7.6. The distribution data of 140 in products
after 2 and 4 min photosynthesis experiments,described in Figure
lO,are presented in the histogram (Figure 11), and may be
directly compared with similar data on the percent distribution
of 14C in products formed by tobacco leaves with and without
OH-PMSulfonate after nearly similar periods of 14002 fixation
(Figure 12). Several differences are apparent. The sugar

lac when the algae

phoSphates (including PGA) contained more
were in OH-PMSulfonate, while in the tobacco leaves with the
sulfonate, the sugar phosphates were lowered; similar changes
were noted in ribulose-l,5-diph08phate.

Most dramatic were the differences in the amount of glyco-
late-140 formation: in the higher plant a tremendous
accumulation of the acid was observed as a result of OH-PMSulfonate
addition, but even at these relatively long times, no similar
effect was shown with the algae. This difference is visually

apparent in the radioautographs from tissue treated with

Figure 8
Radioautographs of Methanol-Water Extract ffiom Tobacco Leaves

After 30 and 300 Second Photosynthesis in 002 in Full Sun-
light

A portion of the percent distribution data in Tables 1

and 2 of the Appendix were obtained from these chromatograms.
Control (Pretreatment 1 hr with water).

Inhibitor (Pretreatment 1 hr with 0.01 M OH-
PMSulfonate).

80

Figure 9

GLYCCLATE

 

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82

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83

Figure 10
Togal Fixation of 14002 by Chlamydomonas reinhardtii at pH
2! _
A l % algal suspension at pH 7.6 in 0.001 M
phosphate buffer. Preincubation time was 5 min in
the light with and without 0.001 M OH-PMSulfonate.

Closed symbols - Control (0.001 M phOSphate)
Open symbols - Inhibitor (0.001 M OH—PMSulfonate)

84

Figure 10

Chlamydomonas (pH 7.6)

(+) OH-PMSulfonate

 

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O
6

804

fi _ _ T
O O
2

.4
Ammmam Ha\muoa H a\ov

aoapomam mandaom ampmzlaoquPoa ad coNHM o Hmpoe

wp

85

mosm ea oowam and .-
oaneoeaamzmumo a Hoo.o . aoaaaaaaH a;
8.5 ma no opoaanoaa a aoo.o . Hoaeaoo ma

 

moo H ad mamoSpGam
nonozm wsaasn_mmsoaovmsmflso an doaaom soapowam oHDSHom Hope:

uHosmSpoz on» ad mposooam osom 9H 03H mo soapsndapmaa psooaom

 

HH ohnwam

86

mQHod 08mm

mnHo4 oszd

oaaa

ma¢noowqw

PP oasmfim

adflRmomm
mdwbm

 

noticed; etqntos laden-Ioueqqem u: pextg otl queOJeg
87

.mopwsmmosaHQ Howsm CH HooNHm 03H I

.m.m ma an opoaoeHamzm
-mo 2 Ho.o aha: as H paoapooaaoam - aoaaaaaaH

 

.aopms SpHs a: H psoapooapoam I Hoapsoo flfl_

 

NODH sH mHmoSpnhm
uoposm wsHasm wobmoq ooomnoe an doahom soHpowaw oHpSHom nope;

IHonozpoz on» :H mpodvoam oaom 3H 03H mo QOHpspHapmHQ psooaom

 

NH oaawam

88

wQHo< OBMM

mQHo4 OZHS¢

oaaa

MBHHooMHU

m_ earmae

mmadmmmomm
m¢wbm

 

noticed; etqntos JadeM-Ioueqiem u: pexp; Otl queoxeg

89

90

OH-PMSulfonate, which indicate the large accumulation of glycolate-
1“C after 30 seconds, as well 300 seconds (Figure 8), while no
increase in glycolate-luc was observed in Chlamydomonas even after
60 seconds photosynthesis (Figure 9).-

A significant inhibition of labeled amino acid formation
probably resulted from a block in the glycolate pathway in the
tobacco experiments; in algae, the sulfonate apparently blocked
the sites of transamination or perhaps glutamate dehydrogenase.
This conclusion is supported by the large decrease of mainly
glycine-17C and serine-14C formed by tobacco leaves with OH-
PMSulfonate, and the accumulation of glycolate-Inc. In the algae,
however, in the presence of the sulfonate the percent 14C in
alanine, aSpartate, and glutamate was more dramatically reduced
than that in serine and glycine. The OH—PMSulfonate in algae
also caused an increased accumulation of 140 in the keto acids,
pyruvate andcx-ketoglutarate; this effect was not observed in
tobacco.

It can be concluded from the above data that the "Specific"
glycolate oxidase inhibitor described by Zelitch (134,137)
functions at sites other than the oxidase. These data from
Chlamydomonas, showing no glycolate accumulation in the presence
of OH-PMSulfonate, can be interpreted to imply that glycolate
oxidase and the glycolate pathway are absent in these algae.

A more thorough investigation of the effects of OH-PMSulfonate

on algae during shorter periods of photosynthesis in bicarbonate

91

was undertaken. The results of these experiments are summarized
under four general headings: (a) the effect of sulfonates on
total fixation; (b) the effect of sulfonates on glycolate
formation; (c) the effect of sulfonates on amino acid formation;
(d) the effect of sulfonates on the sugar diphosphates.

(a) In the presence of the OH-PMSulfonate, 14002 fixation
during photosynthesis is rapid and slightly inhibited at pH 6.5
(Figure 13). On the other hand, the rate of 002 fixation is
normally reduced at pH 8.3 (81), but the addition of OH-
PMSulfonate to the Chlamydomonas stimulated the rate of €02
fixation to that which existed at pH 6.5 even though the pH was
maintained at 8.3 (Figure 14). The reason for stimulation is
not known.

(b) There was no accumulation of glycolate—Inc in the
presence of the OH-PMSulfonate (Figures 15 and 16) at pH 6.5.
or even at the alkaline pH of 8.3 which favors glycolate formation
in algae (81). These results are to be compared with the previous
data from tobacco leaves where the addition of the sulfonate
resulted in the accumulation of large amounts of glycolate.

(c) lb’C incorporation into the amino acids was severely
inhibited by OH-PMSulfonate at either pH 6.5 or 8.3. Correspond-
ingly 11"C labeled keto acids appeared on the chromatograms,
although these keto acids normally do not accumulate in sufficient

amounts to be readily detectable. Thus, in the presence of the

Figure 13
Total Fixation of 14002 by Chlamydombnas reinhardtii at pH
6.5.
Standard photosynthesis experiment with 1 Z algae sus-
pended in 0.001 M phosphate at pH 6.5. Preincubation time
was 5 min in the light with and without 0.001 M OH-

PMSulfonate.

Closed symbols - Control (0.001 M phosphate)
Open symbols - Inhibitor (0.001 M OHHPMSulfonate)

92

Figure 13

Chlamydomonas (pH 6.5)

  

Control
(+) OH—PMSulfonate

  

  

30 60
Time (sec)

20

1O

 

 

—4

160

O“ _

—A

O _ _
2

so
1Ammem Ha\¢roF N a\ov
HOHpowam oHQSHom popoanosospoa sH coNHm UHF

Hosea

93

Figure 14

 

Total Fixation of 11"'C02 by Chlamydomonas reinhardtii
at pH 8.3

Standard photosynthesis epxeriment with l % algae
suspended in 0.001 M phosphate at pH 8.3. Preincu-
bation time was 5 min in the light with and without
0.001 M OH—PMSulfonate.

Closed symbols - Control (0.001 M phosphate)

Open symbols - Inhibitor (0.001 M OH-
PMSulfonate)

94

Figure 14

Chlamydomonas pH 8.3

  

(+) OH-PMSulfonate

 

Control

.1

 

 

O. _

0

Ammem m9\ :0? N S\ov
sOHpowam medHom ampmano waves sH emNHm o

15Q.
_(

wp

proe

6O

3O

20

10

Time (sec)

95

.Aoaosoeasmzmumo a Hoo.ov aopaaassH . nHoaaan soao
.Aopnsanosa z Hoo.ov Hoapsoo u nHonaan eonoHo
.NooaH sH m.m an no pawsHaoawe mHmoanamopona pudendum

 

m.m mAIpm mmsoaouaamHso HQ doaaom
mcHo4 onHad use .opmHooan .dom sH 03H no sOHpanapmHn psooaom

Wfl.oyswdm

 

96

ow om 0% 0%

Aommv oSHB

om om om 9

no an cm 0.

 

 

onesoeasmzaamo A+

   

Hoapcoo

mQHod oszd

 

    

I
‘ 1‘

\

mszmm é mzHowHw unuunuh

QIIO

ma<floowgw

oaneoeasmza-mo A+v

 

  

Hoppcoo

  
  

MadmmOMHwommmomm

I

I

O

I

r

C)
noticed; stqd¥os laden-Ioueqqsm fii pexp; OVI iueoxeg

r

 

 

m_ magma

97

AoeosoeHsmzmumo z Hoo.ov aopapassa . nHonaaw some
Aesosanosa a Hoo.ov Hoaasoo . nHoasan eonoao

.mOUéH :H m.m ma pm psosHaoaHo mHmonpsamoposa pudendum

 

mww mm pm mososouaasto hm weapon
noao< osaaw can .oaoHooaHo .dom ea osH eo soapspaapmao paooaom

 

mH 9dewm

98

00 on om OP

_ b _ L

Aoomv osHe
cm on 0m 0—

om om ON 0—

 

 

     
     

oaesoeaamzaumo A+

 

Hoapcou

mQHod oszd

 

 

Hoapcoo - L

.’ ||||||||||||

33.5% a. mzHoMH cilia:-
mednoowqwollllle

 

Hoapcoo

 

opeCOMHfimzmamo A+v

wadmmownwommmomm

OI

noticed; etqntos laden-Ioueqiem up pext; Otl iueoxea

 

 

o_ oasmaa

99

100

sulfonate inhibitor, 1"(C in alanine was much less, while pyruvate
was strongly labeled; similarly, glutamate-17C was labeled less
anch-ketoglutarate-luc more.

(d) A large accumulation of sugar diphoSphates occurred in
the presence of OH-PMSulfonate (Figure 17). This unusually large
amount of diphOSphate accumulated at both pH 6.5 and pH 8.3.
PhoSphatase hydrolysis of these sugar diphosphate areas from
the chromatograms and rechromatography indicated that the radio-
active material was predominantly ribulose-l,5-diphosphate.

Since the results with Chlamydomonas and Chlorella (data
for Chlorella not presented) in the presence of OH-PMSulfonate
were both unexpected and striking, they were further extended by
repeating similar experiments with hydroxymethanesulfonate
(OH-MSulfonate) another of the sulfonate derivatives which in-
hibits glycolate oxidase (134). The OH-MSulfonate derivative was
chosen because it was not a pyridine analog and its use might
have differentiated between effects caused by the pyridine moiety
and those caused by the<X-hydroxysulfonate moiety.

Results with OHaMSulfonate are presented in Figures 18
through 22 and are essentially similar to the four main effects
on algal metabolism obtained with OH-PMSulfonate. (a) This OH-
MSulfonate apparently had no effect at pH 6.5 (Figure 18), but
stimulated COg fixation at pH 8.1 (Figure 19). This stimulation
restored the rate of fixation at pH 8.1 to that which normally

occurs at pH 6.5. (b) There was no accumulation of glycolate-Inc

Aoeosoeasmzmumo z Hoo.ov aoaaaaasH u mHonsan soao
Aosnsanosa z Hoo.ov Hoapsoo u nHoasan oonoHo

N003H SH m.m no m.m ma pm wusoaHaoaHo meogpsamopona pudendum

 

mmsoaodNSmHso Hm weapon endgamoschun.HuomoHdnHm 2H 03H Hence

 

AH oHdem

101

Om—

. Aommv oaHe

on ON 0—
_ _

omF
_

ow

_ _

Om 0m 0—

_

 

 

Houpcoo

32839.38 3 o

   
  

 
 

m.w ma

 

Hoausoo

opwsoaadmzmumo A+v

m

00 mg

  

I

I

O
dGnH U1 DWI 18101

I

I

C)
a:
(89818 Im/W’Ol x m/o)

 

 

t 8.3m;

102

Figure 18
Total Fixation of 1("C02 by Chlamydomonas reinhardtii at
EH 6.5
Standard photosynthesis experiment with 1 % algae sus-
pended in 0.001 M phOSphate at pH 6.5. Preincubation time
was 5 min in the light with and without 0.001 M OH-methane—

sulfonate.

Closed symbols - Control (0.001 M phosphate)
Open symbols - Inhibitor (0.001 M OH—MSulfonate)

103

Figure 18

Chlamydomonas (pH 6.5)

(+) OH-MSulfonate

Contro

60

20

10

 

 

Q
0

Ammew.wa\
QOHpomam oszHom nopmzuHo

150

m

0.
:J

IOF x B\ov
mapos :H umme o

#—

Hosea

Time (sec)

104

Figure 19

Total Fixation of 14002 by Chlamydomonas reinhardtii at
pH 8.1

Standard photosynthesis experiment with 1 % algae
suspended in 0.001 M phOSphate at pH 8.1. Preincubation
time was 5 min in the light with and without 0.001 M OH-

methanesulfonate.

Closed symbols - Control (0.001 M PhOSphate)
Open symbols - Inhibitor (0.001 M OHHMSulfonate)

105

Figure 19

(+) OH-MSulfonate

 

 

Control

 

 

am

H

P

S

a

n

O

m

0

d

m.

Hm

C
CH 0% C—
a) no .5

AommHm1Ha\ 10* M a\ov

aoHpomaw oHQSHom Hopwleo

m

seems 8a stand 0

qr

anoa

60

3O

2O

10

Time (sec)

106

107

at either pH (Figures 20 and 21). Again this observation stands
in direct opposition to the data expected if glycolate oxidase
were inhibited. (c) The inhibition of amino acid formation was
more pronounced at pH 8.1 than at pH 6.5 (Figures 20 and 21), and
this inhibition suggested that the pyridine moiety of OH—
PMSulfonate was not primarily reSponsible for this effect. (d) A
linear accumulation of 140 in ribulose-1,5-diphosphate occurred
in the presence of the OH-MSulfonate (Figure 22); thus,
reinforcing the similar observation of OH—PMSulfonate's effect

on algae.

Additional data demonstrating the unusual accumulation of
ribulose-1,5-diphosphate-1uc is presented in Figure 23: The
linear accumulation is shown to be a function of the high con-
stant percent 170 in the labeled diphOSphate and the increasing
total fixation of 1["002 by Chlamydomonas. This abnormally high
percent of luc in ribulose-1,5—diphosPhate and the sugar
phosphates in general (Figure 11), and the decreased rate at which
the percent of luc of 3-phosphoglycerate-lnc decreases at pH 6.5
(Figures 15 and 21) suggest that a site ofcx-hydroxysulfonate
activity might exist within the reactions of the sugar phosphates.
Much additional information and data, however, are required
before confirming any relationships or interactions of these
sulfonates and sugar phOSphate metabolism.

Perhaps ribulose-1,5-diphosphate is involved in some reaction

other than 002 fixation, for example, bicarbonate or 002 membrane

RoassocHsmsumo a Hoo.ov aoeanassH : nHoaaan some
Aoaosanosa : Hoo.ov Hoaasoo . nHoaaan eonoHo

.Noo 2H m.w mm as pnoaHHoaHo uHmosusamoposa pudendum

:H

 

I41

. n.w mm1uo mMGOSOUasoHso Np poaaom moHod
osHad use opmHooAHo .dom 2H 03H mo soHpanaumHQ unooaom

 

ON oHSme

108

Aommv maHe

 

 

 

  
 

 

 

          

00 on ON op 00 on ON 0— om on ON or
_ _ _ a _ L _ . _ _ _
QIIIIIIIIIQIHHUQ
opacomgmzlmo T; o a! I Ho.fit.8\0\ -\ 1 \ m 1 \-
Q:\\\\
0 O
Hoapsoo
Hoapsoo
O
onerMHdmzumo A+v
I mszmw e mzHoMHw uIIIIi
mnHo¢ ozH2¢ mawqoownw.?lllllo madmmowqwommwomm

I

O

0' or
w\ m

uotqosx; qunIos Isiah-Iousqiem u: pext; 017l queoxeg

Ci
<1-

 

 

om oasmaa

109

AnassoeHsmzumo a Hoo.ov sopapassa . nHonsan some
Aoadsanosa a Hoo.ov Hoapsoo 1 nHonsan oonoHo
.moodH 2H H.@ mm pm psoaHaoaHo mHmoSpsawoposa camosopm

 

H.m mmInm mmsoaovastno an poahom
moHod osHa< one .opoHooaHo .4om SH 02H no soHpanapmHn psooaom

 

Hm ohszm

110

Aoomv maHs

 

 

on on ON 0— om on ON 0P Om Om ON 0—
r . _ _ _ a L _ a _ _ L
OII O. 111111111 lullulu/

 

38889.3 3

Hoppsoo

mQH04 oszd

 

VIII-.-

mszmM a mzHoMHo
madqoowau

 

 

     

Hoausoo

38882-8 3

mafimmowqwommmomm

 

 

Pm onsmHm

lo 0
CU '—
uotioexg etqntos modem-toueqiem u: pext; on iueoxeg

O
M

O
4-

111

ApposoeHsmaumo z Hoo.ov nopapassH . nHonsan some
Aopmsamosa z Hoo.ov Hospsoo I mHonEam oomoHo

.Noo SH m.m no m.m ma pm mpsoaHaoaNo mHmoSanm090£a pudendum

3H

 

mosoaooaamHno an coaaom opmsmwosmHonm.HlomoHanm :H 03H proa

 

NN onme

112

Om?

om on ON OH

L I

—

Aoomv maHa

F _

ONF

_

cm on ON 0—

. b _

 

 

.I\\\.\.\.

Hoppsoo

onesoeHsmz-ro A+v

   
  

_.m ma

 

HoppCoo

onesoeHsmzomo A+V

m.m ma

 
    

I

r

O

T

O]
(\J
(88319 TW/W_Ol x m/o) dang at 071 18101

 

 

mm

madem

113

.mnsm sH soHpowam HoposlHocmspos on» no 0:H Hence

.mnfim 2H

noHpomam nonexiHosmnpoa can no 0 on» mo psooaom lllll

:H
.AopmsoMHdmzmnmo z Hoo.ov HOpHpHssH I methm cmmoHo
.Aopmsamona z Hoo.ov Hoansoo I mHonahm ammo

.Noo H as n no
no mucoaHaoaNo mHmo£pnmmOpo£m daocsmpm

 

mesoaodNHoHQUANm
doaaoh opmsamosaHdum.Hnom0Hanm :H 03H Hence one usooaom

 

MN oaswam

114

paxu +713 10101 10 °/,

303 08;

ON. 00 On ON 0_

 

r a

IIIIIIIII H. II '0'.

ON .. 282...... s. .00 ..

.93 *0 o\o o

 

808:8 s. 5.0
5:; Ego

m In .6 0088026520 .3
cozoeno... 205000.35 Guam

mm 0.7.1:

2002.30 o: w
o
o_ I \
O

 

O

OO_

OON

00.9

115

some |u1 18d 2_Q| x wdo

116

transport. The stimulated accumulation of ribulose-1,5-
diphosphate-lnC at pH 8.3 was greater than the stimulation
at pH 6.5 (Figures 17 and 22); thus, it is suggested that the
excess sugar diphosphate might account for the increased total
fixation observed at pH 8 (Figures 14 and 19). Further in-
vestigations, perhaps at various C02 concentrations are needed
to completely evaluate this hypothesis.

Asada,§t_§l;.have reported that another of the hydroxy-
sulfonates, Na-glyoxal-bisulfite, reduced the total C02 fixa-
tion by Chlorella ellipsoidea by 50 % (4). Besides the use

 

of a different hydroxysulfonate, they also used a much higher
concentration of bicarbonate (100 uM/3 ml) than was used in my
experiments (less than 10 pM/15 ml). However, the differences
in results and conditions merit further investigations.

The reasons for and the significance of, the four major
observations on the effect of OH—PMSulfonate and OH-MSulfonate
upon algal photosynthesis are not obvious. The inhibitor was
used in all the algae experiments above pH 6, but the tobacco
leaves transpired while in the 0.01 M inhibitor solution at pH
3.5. Since Zelitch has shown up to 60 Z inhibition of glycolate
oxidase at pH 8.3 (134), and that most likely the sulfonate
within the plant cell was no longer at pH 3.5, the block in the
glycolate pathway, caused bycx-hydroxysulfonates will be observed
at physiological pH values (4,137,139). Therefore the differences

117

between the effects of these inhibitors on tobacco and algae
cannot be explained in terms of the pH difference of the¢X-
hydroxysulfonate solutions.

The observed decrease in the amount of glycolate-17C most
likely reflects a block in its synthesis rather than an enhance-
ment of its metabolism. This block in glycolate synthesis might
be related to the accumulation of ribulose-1,5-diphosphate, since
cleavage and oxidation of this diphOSphate between carbons 2 and
3 could lead to phoSphoglycolate formation. No phoSphoglycolate
was observed in any of our experiments with tobacco or algae,
but it has been detected in both plants (11,137). Inefficient
inactivation of the large excess of phosphoglycolate phoSphatase
(92,131) by methanol solutions has been reported by Ullrich (116).
That little or no phosphoglycolate was observed in our experi-
ments probably reflected the poor inactivation of this
phosphatase.

The inhibition of amino acid formation during photosynthesis
by the sulfonate derivatives should be further investigated.

The inhibitor of B6 (pyridoxal phOSphate) transaminases by OH-
PMSulfonate might be expected since both contain the pyridine
moiety. However, the main reason for the inhibition of amino acid
synthesis cannot be explained this way, since the OH-MSulfonate,
containing no pyridine ring, was a very effective inhibitor of

amino acid formation.

118

All the experiments, with both enzymes and inhibitors,
provided no evidence that a glycolate oxidase, having any of the
prOperties of the oxidase of higher plants, was present in algae.
Tanner and Beevers (104) have just published the assumption that
glycolate oxidase "present in algae" was responsible for the in-
creased respiration rates in Chlorella which they observed in
the presence of OH-PMSulfonate. Their assumption is clearly in
error according to the data in this thesis, as well as the data
showing this compound to be an inhibitor of glycolate dependent
02 uptake (137). The observation, however, of increased
respiration in algae merits further investigation before we can
understand the mechanism of OHAPMSulfonate action.

The data presented here might also be compared with the work
of Whittingham's group with Chlorella. In their work, iso-
nicotinyl hydrazide caused glycolate accumulation in algae (84,85).
They interpreted these results as a block in the glycolate pathway
which prevents the conversion of glycine to serine. Yet they
simultaneously observed increased sucrose production. These re—
sults can be better interpreted to suggest that the glycolate
pathway in algae does not contribute to the synthesis of sucrose
as it does in the higher plant. If this pathway were
functioning, then the block in the pathway would cause an increase
in glycolate (as was found withcx-hydroxysulfonate inhibition in
tobacco), but then the end-product, hexoses, should decrease

(Figure 1). The simultaneous accumulation of 14C in sucrose and

119

glycolate by Chlorella in the presence of the inhibitor supports
an argument for a second autonomous pathway for sucrose synthesis
unrelated to glycolate, and that both compounds are likely end—
products of metabolism (125).

Asada,_e_1_: 3f)reported that inhibition of photosynthesis in
spinach leaves by isonicotinylhydrazide caused the expected
accumulation of 1I'I'C in glycine, but none in glycolate. These
data are consistent with the difference between algae and higher
plant metabolism that I have found. In the higher plant
glycolate does not accumulate, although the hydrazide might en-
hance its synthesis; because glycolate oxidase, not inhibited by
isonicotinylhydrazide, causes the conversion of glycolate to
glyoxylate to glycine. Alternatively, because algae lack a
glycolate oxidase these latter reactions are impossible with the
end result that glycolate accumulates.

There are two basic conclusions from these experiments.
First, thecX-hydroxysulfonate inhibitors are not specific for
glycolate oxidase as stated previously. Their apparent
specificity, however, in the higher plant might reflect the large
amount of the oxidase present. Limited absorption and transport
within higher plant tissue and the possible priority given to the
use of thecx-hydroxysulfonates for oxidase inhibition may prevent
achieving concentrations sufficient for effecting amino acid
labeling or for entering the chlor0p1ast to alter sugar phosphate

metabolism.

120

Second, and most important, a comparison of algal and
higher plant metabolism as develOped from these enzymatic, pro-
duct degradation, and in'zixg inhibition studies, reveals that
algae do not have an active glycolate pathway which is typical
of the higher plant. The algae, however, rapidly form and
excrete glycolate during photosynthesis.

Thus, many unsolved problems remain in our understanding of
the metabolic role that glycolate plays in algae and higher
plant metabolism. If the function of glycolate in the higher
higher plant is simply to move carbon from the chlor0p1ast to
the cytoplasm (106,107), then the reason for the excretion of
glycolate to the medium by algae remains obscure. Glycolate
excretion probably represents a more basic function necessary for
maintaining the integrity of the algal cell or the chlorOplast,
for example, ion transport (107).

The difference in glycolate metabolism between algae and
higher plants is also emphasized by the source of glycine in
these organisms. The glycolate pathway in higher plants
adequately accounts for sufficient quantities of uniformly labeled
glycine (107). In algae, however, neither serine nor glycolate
are likely candidates for glycine precursors, since serine does
not account for the labeling pattern in glycine, and glycolate is
not likely oxidized to glyoxylate by glycolate oxidase. Although
Whittingham argues that the effect of isonicotinylhydrazide

121

inhibits the glycine-serine reaction as a B6 inhibitor (89,
124,125), he never considers that other B6 requiring enzymes
like transaminases for glycine (22) should also be inhibited.
Perhaps glycine formation in algae proceeds via some yet

unknown mechanism.

Environmental Factors Affecting Glycolate Production and
Utilization by Algae:

The following series of experiments demonstrated that
pigment changes and metabolic changes did occur when algae
were cultured in limited regions of the Spectrum, i.e.,
different light qualities. The results which support this
hypothesis were obtained from measurements of growth rates,
pigment absorption changes, and the percent distribution of
lLIC in products formed during short time photosynthesis in
various light qualities at various intensities.

Algae cultures, when removed from white light and put in
predominately red or blue light (as described in the methods
section) grew more slowly for 4 or 5 days before attaining a
relatively constant, rapid growth rate which approximated that
of the cultures in white light. Thus, Chlamydomonas and
Chlorella in blue light grew slowly at first and then more
rapidly, as measured by the culture's Absorbance at 680 mu

(Figures 24 and 25). All measurements were made on cultures

which initially had similar cell pOpulations as Judged from

Figure 24

 

Growth Rates of a Chlamydomonas reinhardtii Culture in

Blue Light (400-500 mp)

 

Curve 1:
Curve 2:
Curve 3:

Curve 4:

First day of growth in blue light.
Second day of growth in blue light.
Fifth day of growth in blue light.

Average rate of growth after 10 days
culture in blue light.

122

Absorbance at 680 mu

0
.
14>

Figure 24

 

 

Chlamydomonas reinhardtii

 

Time (hours)

123

 

Figureggj

Growth Rates of a Chlorella pyrenoidosa Culture in

Brie—Eight 1400-500 mp)

 

Curve
Curve
Curve
Curve

Curve

Ut-C'mNH

First day of growth in blue light.
Second day of growth in blue light.
Third day of growth in blue light.
Fourth day of growth in blue light.

Average rate of growth after 10 days
culture in blue light.

124

Figure 25

 

)1

l

O
4:-

l

Absorbance at 680 m

l

 

<1—

 

Chlorella ngenoidosa (Chick)

 

Time (hours)

125

 

126

approximately similar Absorbance values. Chlamydomonas
grown in red light also grew more slowly at first, but after
several days in the red light they also grew more rapidly.
During the first six days of culture in blue light
there appeared a significant decrease in the chlorophyll a/b
ratio when compared to the constant chlorOphyll a/b ratio
from cultures grown in white light. The values reported in
Tablele and 13 are averages of two methanol extractions
from the algae. Variation in total chlorOphyll may be unex—
pectedly large because of the difficulty in accurately
estimating the packed cell volume while using a constant for
this volume in the calculations. The data in Table 12 were
obtained from a different culture than those of Table 13, but
the causes of the large difference in the absolute values of
chlorOphyll a/b ratios and total chlorOphyll remain obscure.
A consistent trend in these data, however, was the decrease
in the chlorOphyll a/b ratio during culture of the algae in
blue light. The data from Chlamydomonas extracts were more
consistent than data from Chlorella, perhaps, because of the
greater difficulty of quantitatively extracting pigments from
Chlorella. To emphasize the general reduction of the
chlorophyll a/b ratio, these data are plotted in Figure 26.
This decrease in the chlorophyll a/b ratio could also be

demonstrated by measurements on whole algae suspensions

Table 12: Chlorophyll Content of Cultures Grown in Blue Light

Amount of chlorOphyll per ml packed cells after growth
in blue light (400-500 mu).

 

 

 

Chlamydomonas Chlorella

Days of Culture Chloro- Ratio Chloro- Ratio
in Blue Light 52711 312:_ $2711 £133_

0 1.09 0.63 0.70 0.77

l 2.03 0.45 1.39 0.60

2 2.15 0.45 1.50 0.36

3 2.63 0.46 1.03 0.77

4 2.02 0.58 1.15 0.71

5 2.56 0.44 2.13 0.51

*Ratio of chlorOphyll a/chlorOphyll b

127

Table 13: Chlorophyll Content of Cultures Grown in Blue Light

Amount of chlorOphyll er ml packed cells after growth in
blue light (400-500 mpg.

 

 

 

Chlamydomonas Chlorella
Days of Culture Chloro- Ratio Chloro- Ratio
in Blue Light fig§ii alb;_ 32%;; alb:_
6 1.55 2.09 1.32 1.88
7 1.83 1.87 1.64 2.01
8 1.54 2.15 1.83 2.38
9 1.69 1.87 1.94 2.11
10 -- -- 1.35 1.49
11 1.64 1.65 ’ —— ——
12 1.53 1.61 1.15 1.83
13 1.82 1.58 1.20 1.86
Cells Grown in
White Light 1.05 2.93 1.14 2.11

*Ratio of chlorOphyll a/chlorOphyll b

128

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129

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¢ m—
a _ _

—~¢

 

 

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130

131

(Figure 27). Using a Cary 15 recording SpectrOphotometer.
absorption Spectra were obtained according to Shibata's
technique (96) modified as described in the methods section.
The Spectra in Figure 27 do have equal Absorbance values at
550 mp, but, for clarity of presentation, the curves have
been separated in the figure. The absorbance change of the
680 mu/655 mp ratio (in Eigg maxima for chlorOphylls a and b
respectively) shows a significant in zivg decrease in the
chlorOphyll a/b ratio with length of culture in blue light.
No significant changes in these regions of the Spectrum were
observed for algae grown in red light. Other regions of the
Spectrum did reflect absorption changes. but these were not
consistent and too complicated for a careful evaluation by
my techniques.

AS indicated by data in Tables 8 and 9 of the Appendix.
a change in the distribution of the 1“C labeled products of
lucoz fixation was apparent in algae after growth in blue
light. In Chlamydomonas (Table 8. Appendix), the most Signi-
ficant change was related to a large increase in the percent
of 14C accumulated in glycolate. but this change was not
observed until Six days after growth in blue light. In
Chlorella. accumulation of excess 1“C in serine occurred almost

immediately after growth in the blue light was begun (Table 9.

Appendix). At all times a consistently high percent of the

.oapmn n Hahnmonoaso

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own no madam» canonnomno Had

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fswaq mfiHm

 

 

ca nacho adpvnmSSHoH monoaoun

 

r ago mo mnpoomm o>d> SH

 

NM ohzwfim

132

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l

 

:E . <
00» 0mm 00m

Onm

-
m><o o
m><o h

m><o N.
m><o N.

on

 

 

 

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5w mpzwfim

N...

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m..—

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N._

aouoq JOSQV

G/v

 

133

134

lhc was incorporated into serine as compared with the percent
found in the control. and a gradual increase in the amount of
lI‘m-glycolate was observed, but this accumulation was not as

large as that observed in Chlamydomonas.

In conclusion, both the pigment data and the fixation
data (particularly that for Chlamydomonas) support an adapta-
tion phenomenon of growth and photosynthesis to blue light.

The most Significant changes which I have observed were a de-
crease in the chlorophyll a/b ratio and an increase in the
accumulation of 1“C in glycolate or serine during Short
periods of 14002 fixation.

Using fully adapted Chlamydomonas (after at least 10 days
growth in either red or blue light), kinetic photosynthesis
experiments were performed in varying intensities of red light,
blue light, or white light. All of the experiments demonstrated
linear fixation rates (Tables 10. 11. and 12, Appendix). A ‘
significant difference was observed with algae grown in blue
light. These algae rapidly accumulated a large percent of the

d 14

newly fixe C into glycolate during a 10 minute period in

blue light (Figure 28). Concurrently there was a decrease in

lac in TCA cycle acids and the associated

the percent of the
amino acids (mainly malate, aSpartate and glutamate) with re-
Spect to algae grown in red light. Apparently the amount of
14C in the sugar phosphate reservoirs in Chlamydomonas adapted

to blue light was much Smaller than in algae adapted to red

N .oupc com um pnmaa eon
ca codpmuam capoSpchmoponm oozfi .pnwaa.uon ca Szonm mHHoo lllll

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caponpeamoposa moo:H .psmaa mean ea neonw maaco

.mhod Cop pwmoa no soy Ala oom
loodv pSwHH QSHD HO Ala OOQAV DSwHH 60H 0» d®pQQUd 0H03 mHHwU

moo SH mamomvzhm
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mQQOSOUhdeSU hp doahom mposuonm SH 03H no Soapsnaapman pnoonom

 

mm shaman

135

ASHBV mafia
or m _

 

I moHod mHoMU 439

 

 

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ll
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O
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136

137

light, Since the percent 140 in these reservoirs decreased

14C in glycine and serine

more rapidly with time. The percent
did not parallel that in glycolate. but instead remained

rather constant and equal for algae adapted to either red or
blue light; thus, the metabolic pools of glycolate and serine
probably are not directly related.

The data in Figures 29 and 30 Should be compared directly
to emphasize the effect that growth of Chlamydomonas in blue
light has on glycolate production. The total 14C in glyco-
late accumulated during photosynthesis in 110 ft-c blue light
almost equaled that fixed at 1200 ft-c white light by cells
adapted to red light. This labeling of glycolate at low blue
light intensity is in contrast to the high light intensity
normally required for glycolate production. It iS also
apparent in Figure 30. that glycolate accumulation increased
much more rapidly than glycine and serine. which iS again con-
sistent with the hypothesis that serine synthesis iS inde-
pendent of glycolate formation and excretion.

The accumulation of labeled carbon in the TCA cycle acids
by red light adapted Chlamydomonas during photosynthesis in
200 ft-c of red light was Slightly greater than that accumulated
by the blue adapted cells utilizing 100 ft-c white light
(Figure 31). As indicated by Figure 32, the total 1“C accumu-

lated in the sugar phOSphates was lower in the blue-adapted

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Acupc oomav unmaa ands: ea mamospeamoposm mIIIII
Ada oomAu.oupc oomv pswaa vow ea mammnpnamoponm .IIII.

 

moo SH mdwoszmm
uoposm meauso Ala ooowv pswaq emm op empamm< Hapunmsnamn

mosoaoumamaso an ozahom Ugo .ondozHc .opmHooan :« o a dance

‘7‘

 

mm mhdwam

138

Figure 29

 

GLYCINE & SERINE

 

 

 

 

GLYCOLATE

 

 

 

 

 

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(I)
739318 TW/Q-Ol x m/o)
UOIQOBJJ atqntos Isiah-tousqiam up paxty o

139

VI

13101

Time (min)

140

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49.3 8.0.5 new: 333 as mammspimoposm Till.
.3: 8m

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H30 an moanom cum .mnHOhHo .mpdaoogau ca 03H Hopoa

  

 

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Figure 30

 

 

 

10

 

 

 

 

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141

71

13101

Time (min)

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A18 oomuoo:v unwaa 03HD 0:99 OHH ma mamospcmmOposm olllllo

.pswda won on confided mowaw

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.pswaa usage oupc ooa ed mdmonpnamosonm QIIIID
.pswaa ands: also coma ea mammSBzamoposm quill.

 

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Hapewwseaow mwcoaoesamaso an cosmos meao< macho «Us ad o:H Hence

 

Hfi‘ohdwam

142

. Anfiav made

n _ o_
L _ r

h—

 

 

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1—

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19101

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11.3

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Hapdnossaou mocoaodwmdflso an monogamonm Howsm ca 0 dance

3
MW 3&3

 

144

Anfiav mafia

op m OF

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145

146

algae than in the red—adapted cells, and this data is con-
sistent with other reports (36,37). Thus. in both algae
adapted to blue or red light, the sugar phosphates were
labeled; however. the pool Size in the blue-adapted
Chlamydomonas appeared smaller as Shown by the large reduction
of the percent lac in these esters (Figure 28). Perhaps these
esters were rapidly metabolized to glycolate which was con-
tinually increasing. 0n the other hand, the large amounts of
phosphate esters accumulated by the red-adapted cells were
probably more Slowly converted in the red light to TCA cycle
acids.

All of the experiments with white light (Tables 10, ll,
12, Appendix) demonstrated that a full Spectrum of light had
little effect on the percent distribution of 1”C in the pro-
ducts formed by the adapted algae. Thus. algae adapted to blue
light produced about the same percent of lac in products formed
during 1 to 10 minutes of lucoz fixation in white light as in
blue light. These facts supported the hypothesis that the
cultures grown in blue light actually experienced a period of
adaptation which altered the cellular constituents. Reports,
on the red light dependent formation of glyceraldehyde phoSphate
dehydrogenase (66). and on the formation of glycolate oxidase in
light (59,109,111),Suggest that light dependent changes in enzymatic

composition of a cell can occur.

147

A series of experiments, with algae adapted to blue light
were performed at various intensities of blue light. to
evaluate the extent of light saturation of 1“C02 fixation by
the available light. Chlorella grown in white light (Figure
33A) and Chlamydomonas adapted to blue light (Figure 33C)
fixed l”C02 with normal light saturation. However, the fixation
rate with blue-adapted Chlorella has an unusual curve Since
very high levels of fixation occurred at very low light in-
tensities (Figure 33B). and further evaluation of this
phenomenon seemed necessary. The percent distribution of 11"C
after 3 and 10 minute exposures of algae (blue-adapted) to
14002 in blue light is presented in Tables 14-17.

Table 14

Percent Distribution of luC in Products Formed by Chlorella
(Adapted to Blue Light) After 3 and 10 Minutes in the Dark
in

 

 

C02

Time of PS 3 min 10 min
Total Fixed

gzm x 10‘3/ml algae .3.4 4.9

% Fixed in:

Aspartate 52.7 44.4
Glutamate 7.5 9.5
Glutamine 32.3 41.3

Misc. 7.5 4.8

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osan op dopmouo Hapdnonnaoh mdSOSOdhmodno no nacho

 

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.psmpa
mafia: ca axonm «maddosonhm oHHoHOHSU "¢.£mmno

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mucoadnomwo
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mm onswam

148

 

10' PS

PS

~s-

 

Figure 33

10' P
3.,ffL/o/////P

Light Intensity (ft-c x 10'?)

O

 

 

 

 

 

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149

CL x m /o)
H19m U1 PGXIJ 071 19101

Table 15: Percent Distribution of 1“LC Fixed by a 0.5 % Sus-
pension of Chlorella (Grown in White Light) During
Photosynthesis in Blue Lightg(400-500 mu)

Time of PS 3 min. 10 min.
Light Intensity 50 ft-c 400 ft-c 50 ft-c 400 ft-c
Total Fixeg

(c/m x 10‘ /ml algae) 8.1 57.1 22.5 242.7

% Fixed in:

PGA 6.8 6.0 10.1 8.9

RuDP -- 10.3 16.9 15.

Sugar-P 21.1 26.1 18.8 33.
2.
2

 

UDPG -- "'" 209

Sucrose --

 

5
8
2
01 "" 02
4
1
O

Glycine _-
Serine 2.0 1
Glycerate -— -_. -_ -_

1
Glycolate -- 4
4
0

 

Alanine 4.1 2.2 5.8 2.8
Pyruvate -— -- -- _-

ASpartate 0
Malate 3
Glutamate 4
Glutamine 2
aeKG -_
Fumarate —— 2,7 -- --
Succinate _- -- -_ _-
Citrate —_ -- a- --

an:

 

Lipids -_ -_
Misc. 4.8 11.4 ~- 5.3

150

Table 16: Percent Distribution of 140 Fixed by a 0.5 1 Sus-
pension of Chlorella (Adapted to Blue Light)
During Photosynthesis in Blue Light (400—500 mu)_

 

 

 

 

Time of PS 3 min. 10 min.
Light Intensity 50 ft-c “400 ft-c 50 ft-c 400 ft-c
Total Fixed

(o/m x 10‘3/ml algae) 13.9 26.1 68.4 148.5
g—Fixed in:

PGA 24.2 14.4 18.4 7.3
RuDP 6.7 22.6 7.3 19.
Sugar-P 26.1 20.5 27.7 20.9
UDPG 1.7 0.2 0.9 1.3
Sucrose 0.4 0.6 1.9 1.1
Glycolate 1.5 3.5 1.1 4.8
Glycine 2.6 .9 3.4 6.7
Serine 7.6 11.5 1 .4 18.6
Glycerate -- -— -- --
Alanine 1.9 1.4 2.9 2.8
Pyruvate -- -- -- --
Aspartate 18.8 7.8 6.6 4.4
Malate 3.9 4.9 3.9 3.1
Glutamate 0.6 -- 1.2 0.7
Glutamine 1.5 1.6 0.8 0.6
q-KG -_ _- -_ __
Fumarate -- 0.6 0.3 0.5
Succinate -- ~- -- 0.6
Citrate -- -- -- --
Lipids -- -- -- --
Misc. 2.4 3.3 11.0 7.0

151

Table 17: Percent Distribution of 1“o Fixed by a 0.5 % Sus-
pension of Chlamydomonas (Adapted to Blue Light)
During Photosynthesis in Blue Light (400-500 mu)

Time of PS
Light Intensity
Total Fixe%

10 min.
50 ft-c 100 ft-c

3 min.
50 ft-c 100 ft-c

 

 

 

 

(g/m x 105_/m1 algae) 8.8 18.0 41.7 186.2
% Fixed in: ‘

PGA 9.8 9.6 4.5 5.8
RuDP -- 10.4 4.2 5.2
Sugar-P 11.3 18.1 24.2 17.7
UDPG -- 1.9 3.3 0.9
Sucrose 2.8 0.4 1.4 1.7
Glycolate 11.2 12.1 19.7 17.5
G1y01ne ‘- o6 209 109
Serine 7.1 6.2 3.9 4.?
Glycerate -- 1.2 -- 0.2
Alanine 7.1 1.5 1.2 1.9
Pyruvate —- -_ _- _-
ASpartate 14.1 5.8 5.3 3.9
Malate 15.5 18.1 11.3 16.4
Glutamate 11.3 . 2.3 4.3 4.5
Glutamine -- 1.9 0.8 4.3
amKG -- -_ __ _-
Fumarate 2.8 1.5 0.6 0.9
Succinate -- -- 0.4 1.5
Citrate _- -— -- --
Lipids -- __ -- --
MiSC. 7.1 3.5 11.1 11.0

152

153

In the dark, normal Chlorella primarily incorporated 1”G
into aSpartate and glutamine as did Chlorella adapted to blue
light (Table 14), as would be expected from C02 exchange
reactions. The percent radioactivity in glutamine‘decreased
at low light intensity, and the percent 1“’0 in aSpartate de-
creased at higher light intensity (Table 15). Blue-adapted
Chlorella accumulated a much smaller percentage of 180 in
aSpartate even at 50 ft-c. and much more 1“C in PGA and RuDP
(Table 16). The very high fixation of 14002 by the blue-
adapted Chlorella (Figure 33B) at low light intensities was
not an expression of increased aSpartate or glutamate metabolism,

but rather of the increased 1”

C accumulation in components of
the photosynthetic carbon cycle.
The blue-adapted Chlamydomonas accumulated an abnormally

r 14c in glycolate, even at low light in-

high percentage 0
tensity and a relatively low percentage in aSpartate and
glutamate (Table 17). The large amount of 140 in glycolate
could be interpreted as a rapid conversion of the sugar phos-
phate pools into glycolate by blue-adapted Chlamydomonas, and
thus both Chlorella and Chlamydomonas respond somewhat Similarly
to growth in blue light. These changes in 1”c product dis-
tribution from 11+C02 fixation suggested that photosynthesis and

metabolism in algae adapted to blue light were altered. Perhaps

154

a more efficient utilization of the incident light energy could
account for the increased photosynthetic C02 fixation at low
light intensities.

Such data. as presented in this section, invite Specu-
lation that perhaps the two major areas of photosynthesis,
electron transport and carbon dioxide fixation. are so closely
related that complementary changes in both systems compensate
for environmental alterations. These changes would tend to
maximize the efficiency with which an organism would photo-
synthesize and metabolize C02. Results from many further
eXperimentS, however. are needed before any final conclusions
can be made with regard to the physiological Significance of
these changes. Complete interpretation of the results with
light quality must await further basic information on the
reasons for and mechanism of glycolate biosynthesis during

photosynthesis.

Manganese Deficient Culture of Chlorella:*

Cultures of Chlorella pyrenoidosa (Chick) were maintained

 

on the inorganic salt medium of Norris et a1. (75). with and

without MnClg added to the micro-nutrients and with continuous

*WOrk done in conjunction with P. L. Youngblood. NSF
Undergraduate Research Participant in 1964.

' 155

0.2 % C02 aeration. The chemicals used were analyzed for
Mn++ and recrystallized in doubly-distilled deionized water.
The Mn-deficient medium was prepared with redistilled de-
ionized water. Only after 20 days. with nutrient changes
every third day. was a reduction of growth rate observed.
The Slower growth rate observed with this culture could be
restored to the normal growth rate by adding an apprOpriate
amount of MnClZ to the deficient medium (Figure 34). The
"Mnerestored" culture achieved normal growth rates after 24
hours. Since the rate curves were measured on cultures with
Similar initial cell pOpulationS, the Mn-deficient rate
actually indicated a Slower log phase of growth (Figure 34).
The fixation data in Table 15 were obtained from standard
photosynthetic experiments in 14C02 (see methods). The
nearly linear rate of total fixation over the 10 minute
period indicated that the lL"C02 concentration never became
limiting. Glycolate production in the manganese deficient
culture was inhibited but glycine and serine synthesis in-
creased. The increased accumulation of 1“C in glycine and
serine in a system not producing glycolate, supports the con-
cept that glycolate oxidase is absent and glycolate conversion
to serine does not occur. The unknown precursors for
glycolate formation might. however, be diverted toward in-

creased glycine and serine production in the manganese

Figureg34

Growth Rates for Cultures of Chlorella.pyrenoidosa (Chick)
During Growth in White Light.

 

Curve 1: normal Chlorella culture.
Curve 2: Manganese deficient culture.

Curve 3: Manganese deficient culture with
MnC12 added.

156

 

Figure 54

 

 

 

36

24

12

 

 

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fla—

hfifi _ 9.:

no
is 0mm 98 moqmpaommw

Time (hours)

157

 

m.mm w.:a m.mH n.0H v.5 w.m nonpo
s.om m.mH m.ma H.0H m.a m.s osppom
s.m m.b H.a m.m N.m b.m cspoaao
s.m m.m 0.: a.om m.sH o.a opoHooaHe
m.ms o.am a.Ho m.sm H.mb m.pa mupcmam
"ca coups a
m.Hps a.maa m.ss p.HHN o.ma N.Hm Accmao

asxmnoa w s\ov

soapasam Hopoa

.oa .m .H .0H .m .H mm po cede

psoaoppoa-sz

Hoppqoo Hosaoz

mm mo GOdpddnoo

 

 

Axoazov omodaocoahm oaaonoaso an doxam 03H no Soapsnaapman paooaom

ma oHQMB

158

159

deficient algae. These results confirm an earlier report on
the requirement for Mn.‘"+ in glycolate formation during photo-
synthesis by Chlorella (103). Tanner;g£‘§1; (103) also re-
ported the increased amount of 1”C labeling of glycine and

serine by manganese deficient Chlorella.

Glycolate Excretion and Uptake:**

The excretion of glycolate by algae has been well docu-
mented (49.70.87.113) and there iS one report of glycolate
excretion by chloroplasts (106). No metabolic function has
yet been elucidated for glycolate excretion. Tolbert (106),
Whittingham and Pritchard (125), Warburg and Krippahl (121).
and 0rth,§t.§l;_(81) have reported maximum glycolate ex-
cretion at low C02 concentrations(0.l to 0.3 Z). high light
intensity, high 02 and high pH. The excretion of glycolate
was measured in the algal supernatant medium as lac-glycolate
or by the colorimetric procedure of Calkins for glycolate
(18). During the course of my research additional conditions
for glycolate excretion and metabolism were considered in our

research group. and the results are reported in this section.

**Uptake and excretion work done in conjunction with
Fang Hui Liao, Graduate Research Assistant. during 1965.

160

The amount of glycolate excreted by the algae was found
to be greatest in the presence of 0.01 M bicarbonate buffer,

but much less in 0.1 M Ncho3 (Table 19).

Table 12
Glycolate Excretion by Chlorella pyrenoidosa (Chick)

 

Glycolate excreted by 2% Chlorella suSpension in water
during photosynthesis in 3000 ft-c.

 

 

Final RCO; Conc. Glycolate in Supernate
moles/l ug/hr/ml
None 9
0.0001 9
0.001 25
0.01 78
0.1 10

In 0.01 M NaHC03 glycolate excretion was linear during the
first hour. These results seem inconsistent with prior obser-
vations on the requirement for low levels of C02 for maximum
glycolate formation. However. all measurements with C02 were
done at pH.rangeS of 4.5 to 6.5. while the pH of a bicarbonate
solution is about 8.3.

The uptake of glycolate by algae has been generally

measured by the disappearance of glycolate determined by the

161

colorimetric assay (26.27.70). Nalewajko,§t_§1l (74) recently
reported. however. a small uptake of 140 labeled glycolate by
a strain of Chlorella isolated from the Arctic Ocean. However.
we have observed. in many attempts with Chlorella and
Chlamydomonas. no Significant uptake of 14C labeled glycolate
or phosphoglycolate during time periods of 10 to 120 minutes.

Typical results are presented in Table 20. The small amount

Table 20

Uptake of Glycolate-2-140 or Phosphoglycolate-Z-luC by Chlamy-
domonas reinhardtii in White Light.

 

A 3 % Chlamydomonas suspension in 0.04 M phOSphate (initial
pH 4.0) in 3000 ft-c White Light.

 

 

 

 

 

Time (min) Percent 14C in Algae
Glycolate-Z-lnc P-glycolate-2~luc

1 1.1 4.1

3 1.3 1.9

10 2.6 2.0

30 0.5 1.0

 

of activity remaining with the cells did not increase with
time. and thus. could have been caused by surface adsorption
of the labeled acid by the algae or filter. The quantitative

colorimetric measurement of glycolate added to algal cultures

162

also indicated no uptake with reSpect to time (Table 21. part
1). In fact. when bicarbonate buffer was used in the pre-
sence of 10'5 M glycolate. a time-dependent excretion of
glycolate with a linear rate occurred for at least one hour.
Thus. the presence of glycolate in the supernatant fluid did
not inhibit further glycolate excretion (Table 21, part 2).

Table 21

Excretion and Uptake of Glycolate by Chlorella pyrenoidosa
Lghiqki. ‘

Part 1: A 2% Chlorella (grown‘ n 5% C02)SuSpenSion was pre-
pared in water and 10‘ M Na-glycolate.

 

 

 

Time (min) Total Glycolate Net Glycolate Change
99751 ps/ml
0 21795 O
5 26.7 +2
15 22.8 -2
30 19.3 -5

60 25.9 +2

 

Part 2: A 2% Chlorella (grown in 5% CO ) suSpension was pre-
pared in 0.01 M NaHC03 and 10-8 M glycolate.

 

 

 

Time (min) Total Glycolate Net Glycolate Change
nS/fil ps/ml
0 21.7 0
5 25.2 +4
15 30.6 +9
30 38.0 +16

60 60.5 +39

 

163

A significant earlier report in 1950 on glycolate meta-
bolism in algae by Schou.gt.§1& (95) claimed that at pH 2.8
in the light. Scenedesmus metabolized glycolate in a period
of 10 minutes to amino acids and sugar phOSphateS. 0n the
basis of this report and the fact that glycolate is formed
rapidly by algae during photosynthesis. the assumption was es-
tablished that the glycolate pathway was present in algae.
These experiments of Schou,§t_§1&.were repeated by feeding
glycolate-Z—luC to three strains of algae: Chlorella

 

pyrenoidosa (Chick). Chlorella pyrenoidosa (Warburg). and
Scenedesmus obliquus (Gaffron D-3) (the same Scenedesmus used
by Schou). Typical results with Chlorella are Shown in Table
22 and Similar results were obtained with Scenedesmus. The
cells were not filtered from the medium as Schou, gt a}; de-
scribed. so that the percent of glycolate conversion could be
estimated. Very little glycolate metabolism was observed
during the 10 minute photosynthesis experiment with glycolate-
2-luC in the light. These data also demonstrate that meta-
bolism at pH 6.0 was equal to or greater than that at pH 2.3

(also found with Scenedesmus).

Table 22 .
Percent Distribution of 114’C After Feeding Glycolate-Z-luc

Approximately 1 no glycolate-Z-lnc was added to 2% Chlorella
pyrenoidosa suSpensionS at pH 2.8 and pH 6.0 in 0.001 M
phOSphate. After 10 min photosynthesis in 300 ft-c white
light the suspension was killed with MeOH and the percent
distribution determined.

 

 

 

 

Chick strain Warburgwstrain
4Com o___und 28.2.8 p121 M 28.22
Glycolate 97.7 98.7 97.7 99.3
Glycine 0.4 0.1 0.3 . 0.1
Serine 0.6 0.3 0.7 1.0
Misc. 1.3 1.2 1.4 4.6

 

No estimation of the percent of glycolate metabolized by
the Scenedesmus was given by Schou,§2.al&, but only the
percent 170 distributed in glycine. serine, and other labeled
products. On the basis of the amount of glycolate-180 used
by Schou,gt'al;'and the intensity of their radioautographs. it
is likely that we have both observed similar limited metabolism
of glycolate-17C by impure (not bacteriaufree) algal sus-
pensions. Several possible reasons might account for the
limited (2 to 3 %) conversion of glycolate-Z-luc to other pro-
ducts. An impurity in the glycolate-14C. such as glyoxylate,
could have been present which could not be detected by the

164

165

chromatographic procedures employed. but which would be
readily used by plant cells (55). As mentioned above the
algal cultures were not free of bacteria which could
possibly account for some glycolate metabolism. There is
also the possibility that the algae have a very limited
ability to metabolize glycolate to glycine and serine.
particularly Since most of the serine in algae iS formed
directly from PGA. The colorimetric determination of glyco-
late disappearance might also be criticized. Since such a
disappearance need not reflect an uptake of the acid by
algae. Another product. e.g. peroxide. also formed by the
cells could cause the breakdown of glycolate; and such inter-
actions could account for the inconsistent results that have
been observed for glycolate uptake and metabolism.

However. from all of the results presented here. I must
conclude that glycolate metabolism in algae is minimal; none
of these data indicate sufficient glycolate uptake or meta-
bolism to contribute Significantly to the metabolism of the
cell or the utilization of the glycolate formed and excreted
during photosynthesis. Speculation. about the role of glyco-
late as an end-product of algal metabolism emphasizes the
need for determining the biochemical reason for glycolate's

formation. The lack of the typical glycolate pathway in algae

166

dictates that we find an alternate metabolic route for glyco-
late utilization. or else discover the biochemical systems

of the cell. dependent upon glycolate formation; so that the
hypothesis of its excretion as an end-product of algal photo-

synthesis becomes rational.

SUMMARY

The Glycolate Pathway in Tobacco Leaves:

Kinetic studies on the rate of glycolate biosynthesis
by tobacco leaves during 11+C02 photosynthesis were run either
in the presence or in the absence ofcx-hydroxypyridinemethane-
sulfonate. an inhibitor of glycolate oxidase. Although 1""002
fixation was reduced about 50 %. the inhibitor did not affect
the initial rate of 1“'C incorporation into phosphoglycerate
or glycolate. nor the Specific activity of these products.
The rate of glycolate formation was slower than the rate of
phosphoglycerate formation. At time periods of 4 and 11
seconds. phosphoglycerate was predominately carboxyl labeled
and glycolate was uniformly labeled. The Specific activity
of the carboxyl carbon of phosphoglycerate was about 100 fold
higher than the Specific activity of either carbon of glyco-
late. Between 30 to 60 seconds. phosphoglycerate became
uniformly labeled and the Specific activities of the carbon
atoms of phosphoglycerate and glycolate became nearly equal.

The inhibitor blocked the glycolate pathway in yizg. AS
a result in a 5 minute experiment the percent of the total
140 in glycolate rose from 3 % in the control to 50 % in the
inhibited leaf. At the same time the percent 1“a in the pro-
ducts of the glycolate pathway. which were formed from
glycolate (glycine. serine. and sucrose). dropped from 32 % to

10 % of the total. These data emphasize the magnitude of the

167

168

glycolate pathway in higher plants. The results extend the
concept that glycolate iS formed after PGA and from a compo-
nent of the photosynthetic carbon cycle. The results do not
support the hypothesis of a second autonomous carboxylation

reaction in photosynthesis for the production of glycolate.

The Glycolate Pathway in Algae:

Enzymes: No glycolate oxidase activity could be de-
tected in cell extracts from four strains of algae. Mano-
metric. isotopic. and SpectrOphotometric techniques were used
to detect the oxidase activity. Repeated attempts to rupture
cells in buffers either with or without glycolate were
unsuccessful in demonstrating the glycolate dependence of
either oxygen uptake or glyoxylate production. Active iso-
citrate dehydrogenase. NADHzglyoxylate reductase. and phOSpho—
glycolate phosphatase were present in the cell extracts. The
lack of a typical glycolate oxidase. suggests that the
glycolate pathway in algae does not exist as it does in the
higher plant. However. these observations do not exclude an

alternative pathway for glycolate metabolism.

Distribution of 1”C in Carbons of Glycolate
PathwangroductS Formed DuriggluCQZPhotosyntheSis in Algae:
The serine formed by Chlorella after 12 seconds contained 80 %
of the total 1“C in the carboxyl carbon and that formed by

Chlamydomonas contained 70 % of the 1“C in the carboxyl carbon.

169

Thus. serine is most probably formed from PGA and not from the
glycolate pathway. Glycine. however. from the same experiments
was uniformly labeled. These results were compared with the
formation of uniformly labeled serine after 4 to 20 seconds
photosynthesis by soybean leaves. The labeling pattern in
serine formed by algae during photosynthesis in 1”C02 demon-
strated that the complete glycolate pathway does not function

in algae.

14

The Effect ofcx-hydroxysulfonates on C02_Photo-

synthesis by Algae: Preincubating l % suspensions of Chlamy-

 

domonas or Chlorella in 0.001 M phosphate with either 0.001 M
a-hydroxypyridinemethanesulfonate or a‘-hydroxymethanesulfonate
produced four main effects on algal photosynthesis in 114002.

Little or no change in total 14

C02 fixation occurred at pH
6.5. but at pH 8.3 the presence of the sulfonate restored the
fixation rate to that at pH 6.5. This rate represented a 3-
fold increase over the fixation rate in the control. In con-
trast to the gigantic accumulation of glycolate observed in
tobacco in the presence of these inhibitors.no glycolate
accumulated in algae at either pH 6.5 or 8.3. Amino acid
synthesis was greatly inhibited in the presence of either «-
hydroxysulfonate as expressed by a decreased accumulation in
alanine. glutamate and aSpartate. Correspondingly there was an
increued accumulation of 1”C in the keto acids. pyruvate and

a—ketoglutarate. These inhibitors at either pH 6.5 or 8.3

also altered sugar phosphate metabolism.

170

A larger percentage of the total lac accumulated in the sugar
phosphates. particularly in ribulose-1.5-diphosphate. From
these results. it was concluded that thecx-hydroxysulfonates
used in this study are not Specific glycolate oxidase inhibi-
tors. These compounds affected both amino acid and sugar
phosphate metabolism in algae. as well as glycolate oxidase in
tobacco. The lack of a typical glycolate oxidase and the
glycolate pathway in algae was supported by the inability of

these inhibitors to cause an increase in glycolate accumulation.

Growth and Photosynthesis of Algae in Red and Blue
Light: Changes in absorption Spectra and 11‘"C02 fixation
patterns were observed with algae grown in red light C>600 mu)
or in blue light (400-500 mp). When grown in blue light
Chlorella and Chlamydomonas Showed a consistent decrease in
their chlorophyll a/b ratio. The Spectra of both methanol ex-
tracts and in 2119 algal suspensions showed this change. The
algae adapted to blue light accumulated 35 % of the 14C
in glycolate; but a normal accumulation was observed in glycine
and serine (7 %). This enhanced accumulation in glycolate
occurred at 110 ft-c of blue light in contrast to the normal
requirement of high light intensity for glycolate formation.
0n the other hand. Chlamydomonas adapted to red light formed
TCA cycle acids and associated amino acids which contained

40 z of the fixed 1“c, and accumulated little or no lie in

171

glycolate (5 %) during photosynthesis in 200 ft-c of red
light. The percent of sugar phOSphates formed by cells
adapted to red light was greater after 10 minutes (40 %) than
the percent of 1“c in these compounds formed by algae adapted
to blue light (20 %). That glycolate accumulation increased
without much change in the percent 14c in glycine and serine
is consistent with the hypothesis that serine synthesis is
independent of glycolate formation. i.e.. these algae did not
contain a complete glycolate pathway. _
Further experiments with algae adapted to blue light re-
vealed that even at low blue light intensities (50-100 ft-c)
the cells fixed a higher percent of 14C in the sugar phosphates
than did a control culture grown in white light. The white
grown cells formed mainly aSpartate and glutamine from 17002
at this low intensity of blue light. Such experimental results
could suggest that the systems~for 002 fixation and electron
transport are related in their response to environmental
conditions.

Manganese Deficient Culture of Chlorella: Manganese
deficient cultures were obtained after 20 days growth in a Mn++
deficient medium. The decreased growth rate of this culture
could be restored to a normal rate by the addition of MnCl2 to
the culture. After photosynthesis for 10 minutes the percent
14c in glycolate formed by normal algae was 30 % while that
formed by the deficient algae was only 2 %. Concurrently there

172

was an increase in the percent of glycine and serine formed
by the Mn—deficient Chlorella. These data also suggest that
glycolate metabolism to form serine does not occur as it

does in the higher plant.

Glycolate Excretion and Uptake by Algae: The amount
of glycolate uptake by algae was measured either by dis-
appearance of the glycolate in the medium or by appearance of
labeled glycolate or phosphoglycolate in the cells. No time
dependent uptake of Significant amounts of either glycolate or
phosphoglycolate was observed. Feeding glycolate-2-14C to
Chlorella and Scenedesmus revealed that only 2-3 % of the
radioactive acid was metabolized by these cultures after 10
minutes photosynthesis. Several reasons. including
bacterial contamination or impure substrates. could explain
this small percent conversion. Again the conclusion must be
that the algae were not capable of glycolate metabolism as is
the higher plant. Glycolate excretion was maximal in 0.01 M
bicarbonate and occurred in the light even in the presence of

10'5 M glycolate.

Comparison of Glycolate Metabolism in Algae and
Tobacco Leaves: From the results in this thesis it is con-
cluded that there exists a basic difference between the
metabolism of glycolate by algae and that of the higher plant.

In both plant tissues, 3-phosphoglycerate is the initial

173

product of photosynthesis with the subsequent formation of
glycolate from the carbon reduction cycle. However. only in
the higher plant is there glycolate oxidase and the subse-
quent reactions of the glycolate pathway. Algae contain no
typical glycolate oxidase and therefore do not produce
glycine. serine or sugars from glycolate as does the higher
plant. Instead the algae excrete the glycolate into the

medium as an apparent end product of metabolism.

10.

11.

12.

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Properties of a new
Biochem. J. 82:

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